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FLORAL FRAGRANCE, POLLINATION, AND SEED GERMINATION OF TWO NATIVE, EPIPHYTIC ORCHIDS IN SOUTH FLORIDA

HALEIGH AMANDA RAY

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2018

To my family and friends who have been tremendously encouraging

ACKNOWLEDGMENTS

I am extremely grateful for all of my friends and family members who have given endless amounts of love, support, and encouragement as I have progressed through my Ph.D. completion. Without them, this degree would not have been possible.

I would like to thank my committee chair and advisor, Dr. Jennifer L. Gillett-Kaufman, for all of the support and advice she has provided during my time here. Thank you for your patience and motivation throughout my dissertation research, it has helped prepare me for my all of my future work in addition to everything that I have completed here at the University of

Florida.

Additional thanks to my committee members, Dr. Michael Kane, Dr. Charles Stuhl, Dr.

Jaret Daniels, and Dr. Jaime Ellis for their feedback on my research. Dr. Kane was always available to help with my seed germination, from the use of his laboratory to troubleshooting research obstacles. Dr. Stuhl provided tremendous assistance in processing of floral fragrance samples and working with me to fully describe the resulting data.

I am also very appreciative to the members of the Gillett-Kaufman laboratory, including

Dr. Morgan Byron, Eleanor Phillips, Dr. Lawrence Reeves, Omotola Dosunmu, Dr. Eutychus

Kariuki, and Ryan Fessenden for review and editing of my dissertation. Additionally, I would like to thank Dr. Lawrence Zettler and Dr. Marjorie Hoy for all of the guidance and preparation they have given me on the path to my dissertation. Although unnamed here the other faculty and staff at the UF Entomology and Nematology Department have always been supportive of my work and made me feel at home in Gainesville.

This research could not have been carried out without the assistance of the U.S. Fish and

Wildlife staff at the Florida Panther National Wildlife Refuge (FPNWR), especially Mark

Danaher and Ben Nottingham for coordinating use of refuge facilities and monitoring orchid

populations. I am very appreciative of the field work assistance at FPNWR that I received from

Larry Richardson, Andrew Stice, and Fauve Wilson, all of whom spent exhausting hours in the refuge with me collecting data. Additionally, the hours in the laboratory from Brandon Corder assisting in orchid seed counting are much appreciated. I would also like to thank James Colee of the University of Florida IFAS Statistics Department for statistical consulting.

Finally, I would like to thank the University of Florida Graduate School for the Graduate

School Fellowship that I was awarded, allowing me to conduct my dissertation research over the last three and a half years.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 8

LIST OF FIGURES ...... 9

ABSTRACT ...... 11

CHAPTER

1 LITERATURE REVIEW ...... 13

Pollination ...... 13 Florida Ecosystems ...... 14 Insect and Vertebrate Pollinators ...... 16 Hymenoptera ...... 16 Coleoptera ...... 19 Diptera ...... 20 Lepidoptera ...... 23 Birds ...... 25 Bats ...... 27 Orchids and Orchid Pollination ...... 28

2 FLORAL FRAGRANCE ANALYSIS OF Prosthechea cochleata (ORCHIDACEAE), AN ENDANGERED NATIVE, EPIPHYTIC ORCHID, IN FLORIDA ...... 39

Introduction ...... 39 Materials and Methods ...... 40 Fragrance Collection ...... 40 Identification of Floral Odor ...... 42 Results...... 43 Discussion ...... 43

3 POLLINATION OF Encyclia tampensis, THE COMMERCIALLY EXPLOITED BUTTERFLY ORCHID, BY VARIOUS INSECT TAXA IN SOUTH FLORIDA ...... 50

Introduction ...... 50 Materials and Methods ...... 51 Pollinator Exclusion ...... 51 Pollinator Collection ...... 52 Statistical Methods ...... 53 Results...... 53 Pollinator Exclusion ...... 53 Pollinator Collection ...... 54

Discussion ...... 56

4 EFFECT OF POLLINATION AND LOCATION ON SEED GERMINATION OF TWO NATIVE ORCHIDS ...... 67

Introduction ...... 67 Materials and Methods ...... 68 Prosthechea cochleata ...... 68 Encyclia tampensis ...... 70 Statistical Methods ...... 70 Results...... 70 Prosthechea cochleata Seed Capsules ...... 70 Encyclia tampensis Seed Capsules ...... 71 Discussion ...... 72

5 DETERMINING POTENTIAL POLLINATORS OF Prosthechea cochleata (ORCHIDACEAE) AND ATTRACTION OF HONEY BEES TO A P. cochleata SYNTHETIC FLORAL FRAGRANCE BLEND ...... 80

Introduction ...... 80 Methods ...... 81 Pollinator Sampling ...... 81 Floral Odor ...... 82 Insect Trapping ...... 82 Y-tube Behavior ...... 83 Statistics ...... 84 Results...... 85 Pollinator Sampling ...... 85 Insect Trapping ...... 85 Y-tube Behavior ...... 85 Discussion ...... 86

6 CONCLUSIONS AND FUTURE WORK ...... 91

APPENDIX

A INSECTS COLLECTED FROM THE FLORIDA PANTHER NATIONAL WILDLIFE REFUGE DURING Encyclia tampensis BLOOMING ...... 98

B INSECTS COLLECTED FROM TRAPS CONTAINING SYNTHETIC ORCHID FRAGRANCE ...... 104

LIST OF REFERENCES ...... 105

BIOGRAPHICAL SKETCH ...... 118

LIST OF TABLES

Table page

2-1 Sample schedule for each of three Prosthechea cochleata flowers from different plants at the Entomology and Nematology Department (UF), and the single flower at the Florida Panther National Wildlife Refuge (FPNWR)...... 46

2-2 A list of the top eight compounds detected in the floral fragrance samples of Prosthechea cochleata ...... 47

3-1 Three year mean percentage of seed capsule formation (±SD) for flowers at each of the locations where Encyclia tampensis was studied at the Florida Panther National Wildlife Refuge...... 59

3-2 Identified plants that were in bloom at each of the four locations across the Florida Panther National Wildlife Refuge that were within 10 meters of blooming Encyclia tampensis flowers...... 59

5-1 Eight compounds created the floral blend that was in each of the six traps...... 89

A-1 Insects collected from the Florida Panther National Wildlife Refuge during Encyclia tampensis blooming from 2015-2017...... 98

B-1 Insects collected from traps containing synthetic orchid fragrance. Preliminary data collected in 2017...... 104

LIST OF FIGURES

Figure page

1-1 The basic structure of a flower including both male and female reproductive parts...... 34

1-2 Map of Florida representing conservation land and strategic habitat necessary for maintaining long-term biodiversity...... 35

1-3 Diagram highlighting the pollen basket on the leg of a honey bee (Apis mellifera)...... 35

1-4 Dendrophylax lindenii, the ghost orchid, in south Florida. The red arrow is pointing to the flowers nectar spur, which suggests a hawk moth pollinator with a long probosics ...... 36

1-5 Inflorescences of Satyrium coriifolium orchids in bloom...... 37

1-6 Structure of a flower in the family Orchidaceae...... 38

2-1 Flower of Prosthechea cochleata, the clamshell orchid, growing at the Florida Panther National Wildlife Refuge ...... 48

2-2 Image showing the collection of floral volatiles from the greenhouse grown Prosthechea cochleata ...... 49

3-1 Butterfly orchid (Encyclia tampensis) and orchid pollinia...... 60

3-2 Map of the four locations at the Florida Panther National Wildlife Refuge from which Encyclia tampensis flowers were sampled during the three-year study...... 61

3-3 A fine mesh exclusion bag placed around two Encyclia tampensis flowers that had opened soon after being covered...... 62

3-4 Flowers of Encyclia tampensis...... 63

3-5 Three types of insect traps were used to sample potential pollinators of Encyclia tampensis, the Florida butterfly orchid...... 64

3-6 Total number of insects actively sampled from each location at the Florida Panther National Wildlife Refuge where Encyclia tampensis was flowering over a three-year period (2015-2017)...... 65

3-7 Total number of insects in each order actively sampled from Encyclia tampensis flowers at the Florida Panther National Wildlife Refuge over a three-year period (2015-2017)...... 65

3-8 Difference in average number of insects sampled around Encyclia tampensis flowers in the morning (0800-1200 h - AM) compared to the afternoon (1230-1630 h - PM) at four locations in the Florida Panther National Wildlife Refuge...... 66

4-1 Mesh exclusion bag placed on an unopened bloom of Prosthechea cochleata in the Florida Panther National Wildlife Refuge...... 74

4-2 Example of four sub-replicates of Prosthechea cochleata seeds on P723 orchid seed sowing agar Petri dish...... 75

4-3 Percent total germination of Prosthechea cochleata seed capsules covered with exclusion mesh (Bagged) at the Florida Panther National Wildlife Refuge (FPNWR), hand cross-pollinated (Hand Crossed), or left open at the FPNWR (Open)...... 76

4-4 Comparison of Prosthechea cochleata seed germination across three locations in the Florida Panther National Wildlife Refuge (FPNWR)...... 77

4-5 Six Prostheceha cochleata seeds (Stage 5) among numerous other seeds that did not germinate...... 78

4-6 The percentage of germinating Encyclia tampensis seeds from capsules in each of three locations at the Florida Panther National Wildlife Refuge...... 79

5-1 Y-tube olfactometer set up in the laboratory...... 90

6-1 Preliminary results of an ecological web created with the R package 'bipartite' comparing orchid species to their identified pollinator...... 97

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

FLORAL FRAGRANCE, POLLINATION, AND SEED GERMINATION OF TWO NATIVE, EPIPHYTIC ORCHIDS IN SOUTH FLORIDA

Haleigh Amanda Ray

May 2018

Chair: Jennifer L. Gillett-Kaufman Major: Entomology and Nematology

South Florida is home to a number of native orchid species. The Florida Panther National

Wildlife Refuge (FPNWR) in Collier County, Florida, has 27 known species, including

Prosthechea cochleata, the clamshell orchid, and Encyclia tampensis, the Florida butterfly orchid. These orchids are listed as endangered and commercially exploited (respectively) on

Florida's Regulated Plant Index. The goal of this research was to determine aspects of the pollination biology for these two species. Several studies were conducted including floral fragrance volatile collection, potential insect pollinator trapping, pollinator exclusion studies, and asymbiotic germination of resulting seed capsules.

Floral fragrance was collected from P. cochleata flowers both in the greenhouse and at

FPNWR by using GC/MS analysis of headspace volatile collection. The orchids sampled consistently produced eight volatiles that are common in floral fragrances, including those noted previously in other orchid species.

Potential pollinators were collected from areas near flowering orchids at FPNWR using several different methods during the peak flowering time. In addition to over 200 combined hours of active sampling from both E. tampensis and P. cochleata, three types of insect traps were deployed. It was determined that E. tampensis is pollinated by a wide range of insect

orders, while only one Hymenopteran family (Apidae) was actively collected from the flowers during P. cochleata blooming.

Exclusion studies were conducted on greenhouse grown specimens of both orchid species at the University of Florida and on plants growing in the wild at FPNWR by either exposing or covering flowers using exclusion netting to prevent any insect interaction. The covered flowers of E. tampensis had no seed capsule formation, while the flowers of P. cochleata had over 65% formation, suggesting that this species may be capable of autogamy.

Seed capsules produced during this study were collected and taken to the laboratory for asymbiotic germination. Germination rates were compared for E. tampensis capsules produced from different locations at FPNWR, and similarly for P. cochleata, but including capsules that were formed inside or outside of exclusion bags. This research produced promising results that can help develop future conservation efforts to protect and increase populations of these orchids.

CHAPTER 1 LITERATURE REVIEW

Pollination

Plants reproduce sexually by pollen transfer. Each flower can contain male and/or female reproductive structures. The female part of a flower is the pistil, and the male part is the stamen

(Figure 1-1). The pistil is comprised of the ovary, ovule, stigma, and style, while the stamen consists of the filaments and the anthers, which is where the pollen is produced. A plant will produce gametophytes, or the sexual phase, which then produce male and/or female gametes. In gymnosperms and angiosperms, the female gametophyte is formed in the ovule of the flower, and the male gametophyte, the pollen, is formed on and moved from the anther of the flower to the stigma of either the same flower or a different flower of the same species by wind or by a pollinator (Holsinger 2000).

Pollination takes place when the pollen grains are moved from the anthers to a stigma.

Once the pollen comes in contact with the stigma, a pollen tube begins to form (University of

California-Davis 2013). In nature, many species of flowering plants are only able to complete the pollination process with the help of pollinators that transfer pollen between their flowers.

Approximately 80% of flowering plant species depend on animal pollinators, typically insects, most commonly Coleoptera, Diptera, Hymenoptera, and Lepidoptera (Food and Agriculture

Organization of the United Nations 2008). Pollinators are well known for their importance in agriculture and crop production, and are critical to maintain biodiversity, and for supporting populations of native flowers. Maintaining a diverse plant population indirectly affects animal species that are dependent on floral resources, like seed eating animals and herbivores (Potts et al. 2006).

Pollination services are facing increasing pressure from several sources including habitat fragmentation, loss, and isolation, increased monoculture, pesticide use, introduction of non- native species (including pests of hymenoptera), climate change, and competition for resources with managed pollinators (Jennersten 1988, O'Toole 1993, Frankie et al. 1997, Kearns et al.

1998, Price and Waser 1998, Steffan-Dewenter and Tscharntke 1999, Donaldson et al. 2001,

Gross 2001, Brown et al. 2002, Kremen et al. 2002, Battacharya et al. 2003, Thomson 2004).

Despite the importance of pollination, the value of pollinators for biodiversity and plant- pollinator communities is relatively understudied. It is known that some wild plants can have significant pollen limitation (Burd 1994). In a literature review by Morales and Traveset (2009), a negative effect on pollination of native flowering plants that were located near non-native flowering plants was confirmed. This effect was greatest when the non-native species had a similar flower color and symmetry to the native flowering plants (Morales and Traveset 2009).

Williams et al. (2015) tested wildflower mixes planted near crops across the United States to measure the species richness and abundance of pollinators. They found that the mixes increased the richness and abundance of wild bees in most of the ecosystems tested.

Florida Ecosystems

Florida is a biologically diverse state and includes both temperate and sub-tropical plant and animal species. Researchers have referred to the necessity of identifying the habitats that are critical to support both rare and endangered species and other biodiversity factors before they are lost (Noss and Cooperrider 1994). The number of tropical species in Florida increases moving south through the peninsula (Ewel 1990). Like many regions, native Florida habitats are disappearing at a rapid rate. According to Noss and Peters (1995), Florida was ranked as the state with the greatest risk for biodiversity loss. Human disturbance from a growing population is

threatening the survival and sustainability of Florida ecosystems by causing habitat loss and fragmentation (Pearlstine et al. 2002).

Kautz and Cox (2001) identified 40 focal species of plants that needed habitat conservation. They then identified areas of Florida that are conservation land or what they described as strategic habitat, or "land that, if conserved, would have the best chance of enhancing long-term survival potential of the 40 focal species as well as other components of biodiversity in Florida" (Figure 1-2).

Many longleaf pine habitats in the southern United States have been severely changed or lost all together. One study by Pitts-Singer et al. (2002) surveyed longleaf pine habitats in the

Apalachicola National Forest in Florida. They determined potential insect pollinators of three

Florida native flowers located there, but noted that the rate of flower visitation by these insects was very low (Pitts-Singer et al. 2002). This raises concerns about the loss of forest habitat affecting the survival of other native plants and pollinators.

Another study by Liu et al. (2006) compared native and introduced species pollinating an invasive vine, Paederia foetida L. (skunk vine), in Florida. They found that both halictid bees that are native to Florida and the introduced western honey bee (Apis mellifera L.) were pollinating this plant. They also noted that floral visits by the native bee species was much more common in less disturbed sites, while the honey bee visited the plant more frequently in disturbed sites (Liu et al. 2006). This suggests that the native bees could be driven away from disturbed areas, and could become threatened with the increase in urban development across

Florida.

Insect and Vertebrate Pollinators

Hymenoptera

More than three-quarters of all flowering plants must be pollinated by an animal visitor, usually an insect (Klein et al. 2007). It is estimated that 71 of the 100 crop species which provide

90% of food worldwide are bee-pollinated. These food crops include fruit, nut, and seed crops

(United Nations Environmental Programme 2010). It is estimated that one-third of food humans consume comes from plants that are pollinated by honey bees, either directly or indirectly (Free

1993).

Of over 20,000 bee species worldwide, the most recognized of these is the western honey bee, Apis mellifera L. There are several characteristics that make the honey bee a good pollinator. The population in an area can be easily increased by adding hives. Moving hives from one location to another can give the bees a better chance to find and pollinate targeted plants or crops (Jaycox 1976). Honey bees require large amounts of pollen and nectar to rear progeny, making it necessary to visit flowers regularly to attain these food resources. This often leads to the honey bees visiting one species of plant at a time, increasing the likelihood of successful pollen transfer (Jaycox 1976).

The physical characteristics of the honey bee are beneficial to pollination. Like many other bees, A. mellifera bodies have small, branched hairs that help hold pollen that is being carried from one flower to another. Bees often store pollen in a corbicula, or pollen basket on their rear legs (Figure 1-3), and these hairs trap the pollen where it can be inadvertently transferred to another flower visited by the bee. Honey bees have a medium body size, allowing them to pollinate a wide range of plant species with different flower types (Jaycox 1976.)

Globally honey bee populations have been declining in recent years. The decline began in

2006, when beekeepers reported higher than normal colony decline, losing anywhere from 30-

90% of their colonies (United States Department of Agriculture, ARS 2008). The cause of this loss is believed to be a multitude of factors including exposure to pesticides, parasitic mites (such as Varroa destructor Anderson and Truemann), insufficient food supply, and viruses that infect the bees and slow their immune systems (Cox-Foster et al. 2007, Natural Resources Defense

Council 2008, Ellis et al. 2010). Non-Apis bees may be less affected by some of these factors, for example, the parasitic mite V. destructor affects only bees in the genus Apis (Committee on the

Status of Pollinators in North America, National Research Council 2007). Varroa destructor is one of the most serious threats to honey bees. The mites are very small, and feed on the circulatory fluids of the bee, spreading viral and bacterial pathogens that can cause disease

(United Nations Environmental Programme 2010).

The value of crops pollinated by honey bees is over $215 billion worldwide, and greater than $14 million in the United States, putting the U.S. at risk of losing this important contribution to its gross domestic products without honey bees to pollinate the crops (Morse and

Calderone 2000, Natural Resources Defense Council 2008, van Engelsdorp et al. 2008).

There are over 4,000 species of native bees in the United States. Though honey bees are well known, they may not be as probable pollinators as native bees in pollination of native plants. Aside from pollination of crops, native bees are highly valuable to the ecosystem. They are responsible for pollinating up to 80% of flowering plants (Moisset and Buchmann 2011). Of all plants worldwide, it is estimated that about 20% are threatened with extinction. In addition to this, there have been predictions of large, negative changes in the next 50-100 years to all terrestrial plants (Tilman and Lehman 2001). While some of these changes are the result of habitat destruction, other human interference affecting soil composition, an increase of CO2 in the atmosphere, disturbances to populations of animals that are herbivores or predators, and

climate change are cause for concern as well (Tilman and Lehman 2001). Currently, plants are equally as threatened as mammals in terms of extinction risk, and more threatened than birds

(Ibrahim et al. 2013). All species need a range of particular combination of biotic and abiotic conditions to survive, known as a fundamental niche. If these conditions are changed or destroyed, that species is no longer able to survive in its environment.

As many species are at a large risk of extinction, it is important for humans to understand the effect that they have on these species, and the value that these species have, both ecological and economic. Many endangered plant species belong to families that have other species in them that produce already utilized pharmaceutical drugs (Ibrahim et al. 2013). Of products approved by the Food and Drug Administration (FDA) in 2009-2010, 26% of them were, in some way, naturally derived (Zhu et al. 2011). Additionally, in a study by Newman and Cragg (2007) it was estimated that higher than 60% of cancer therapeutics were derived from naturally occurring products. A loss of species richness globally could reduce the opportunities we have to develop new drugs and novel treatments in the future.

The morphology of native bee species is diverse, with many different shapes, colors, and sizes (Moisset and Buchmann 2011). Native bees provide a significant contribution to crop pollination as well as filling an important role in many ecosystems by pollinating wild plants

(Kremen et al. 2002, Winfree et al. 2007). Because other bee species (honey bees and bumble bees) can be rented or purchased by farmers to enhance crop production (Morse and Calderone

2000), it suggests that native bee pollinators alone may not sufficiently pollinate crops that are planted in large monocultures. Winfree et al. (2007) concluded that in 23 watermelon farms throughout the state of New Jersey, native bee species provided over 90% of pollination services at the study sites. Native bees have been documented to pollinate coffee, canola, sunflower, and

other crops (Ricketts 2004, Morandin and Winston 2005, Greenleaf and Kremen 2006, Klein et al. 2007), but previous studies indicate that native bees are insufficient crop pollinators (Kremen et al. 2002). The success of native bees in crop pollination may vary depending on the type of crop, the size of the farm, and the species of bee. Javorek et al. (2002) compared the effectiveness of native and non-native bee pollinators on lowbush blueberry (Ericaceae:

Vaccinium angustifoli). They found that Bombus spp. and Andrena spp. bees pollinated flowers

3.5-6.5 times faster than A. mellifera, and that it would take A. mellifera four visits to a flower to deposit the same amount of pollen that a Bombus spp. and Andrena spp. deposited in one visit.

Honey bees are not the only bees that are facing population challenges. Populations of some native bees are declining due to similar factors that are affecting honey bees. Habitat loss and fragmentation, pesticide use, or fewer nectar and pollen resources are contributing to population declines of native bees (Moisset and Buchmann 2011). Habitat fragmentation causes food sources to become reduced and inadequate to support the pollinator populations. This also results in a loss of habitat for nesting, and foraging (United Nations Environmental Programme

2010). The International Union for Conservation of Nature (IUCN) has predicted a loss of nearly

20,000 species of flowering plants globally within a few decades (United Nations Environmental

Programme 2010). This will likely cause a decline of pollinators that rely on these plants to survive.

Coleoptera

Coleoptera, or beetles, are another important group of insect pollinators. Based on insect fossil records, it is believed that the order Coleoptera were the first insects to be associated with floral visitation (Baker and Hurd 1968). From there, insects from the orders Lepidoptera,

Diptera, and then Hymenoptera started to become connected to flowers as well. According to a study by Thien (1980), this suggests that a significant number of primitive angiosperms were

pollinated by Coleoptera. While some beetle-pollinated flower species are generalist pollinated, meaning they can be pollinated by a wide range of insect orders, others are specialist pollinated, and have an obligatory relationship with a particular beetle (Bernhardt 2000). Typically, most beetle-pollinated flowers are bowl-shaped, large, and have abundant amounts of pollen present.

They may also have large, leaf-like stamens adapted for pollination by beetles (Thien 1980).

While many of the flowers that are beetle-pollinated are wild, some beetles serve as pollinators for commercial crops. In Florida, Annona species were found to only set fruit when visited by beetles in the family Nitidulidae (Nadel and Peña 1994). The beetles enter the flowers and remain there until senescence, and the number of nitidulids correlates with the percentage of flowers that set fruit.

Some flowers in the family Orchidaceae are pollinated by beetles. Most often, this comes in the form of deceptive pollination. Orchids are known for the high number of non-rewarding species in comparison to other types of flowering plants (Jersáková et al. 2006). Some common types of deception in orchids are food deception, sexual deception, and brood-site imitation. In food deception, the pollinator believes it will receive a pollen or nectar reward when one is not actually present. Whereas in sexual deception, the flowers resemble a female insect, and when a male attempts to mate with multiple flowers, pollen is subsequently transferred between them.

Orchids with brood-site imitation attracts insects looking for a place to deposit eggs, for example carrion or fungal fruiting bodies (Jersáková et al. 2006). For beetles, deception typically comes in the form of food rewards or brood-sites, but sexual deception has been documented from several species of Ophrys orchids (Paulus 2006, Tyteca et al. 2006).

Diptera

Along with Hymenoptera and Coleoptera, Diptera, the flies, are another group of important insect pollinators. There are over 70 families that have flower visiting species, which

can be pollinators for more than 550 flower plants (Kearns 2001, Larson et al. 2001). Diptera could have been some of the first pollinators of angiosperms, playing an important role in their evolution (Labandeira 1998, Endress 2001). Like insects of other orders, some Diptera feed on nectar and/or pollen exclusively, and other flies will feed on them opportunistically. Typically specialized pollinators result in higher plant pollination success, but according to some studies, generalist Diptera are significant contributors to pollination success and can be very effective

(Motten 1986, McGuire and Armbruster 1991, Kearns and Inouye 1994).

Diptera are important not only for wild plants, but for cultivated agricultural crops as well, and for enhancing fruit production (Ssymank et al. 2008). Examples of cultivated crops that can be Diptera pollinated are mango, cacao, carrots, apples, and cassava (Hansen 1983, Larson et al. 2001, Mitra and Banerjee 2007). A species from the family Syrphidae has been documented to be a successful pollinator of peppers in a greenhouse setting (Jarlan et al. 1997).

While the most common family of flower visiting flies is the Syrphidae, flies outside of this family have been found to carry 84% of pollen moved by Diptera in farmlands (Orford et al.

2015).

Diptera have been recorded as pollinators of some orchid species as well, even though they are not as common pollinators compared to Hymenoptera and Lepidoptera. van der Pijl

(1961) described the labellum of some orchids as tools used to direct some fly species to the stigma of their flower, in order for pollination to occur successfully. While some of the flowers are visited by flies for a nectar or pollen reward, some flowers attract flies by producing visual or olfactory cues (Ssymank et al. 2008).

One example is the terrestrial orchid Epipactis veratrifolia, which is pollinated by flower flies in the family Syrphidae (also known as hoverflies). The larvae of the hoverfly species that

visit E. veratrifolia are not very mobile after egg hatch, and they feed on aphids while the adults are nectar feeders (Stökl et al. 2010). These orchid flowers produce a fragrance that mimics alarm pheromones of aphids. Because the hoverfly larvae do not disperse long distances, the adult female will search for a place to lay her eggs near an aphid food source, or when attracted to the mimic alarm pheromone inside the flower, coming in contact with the pollinia (Stökl et al.

2010). She will then often move to other flowers searching for a good location to deposit additional eggs, resulting in pollination of E. veratrifolia.

Hoverfly pollination of orchids is not limited to mimicking prey pheromones. A lady slipper orchid (Paphiopedilum barbigerum) is visited by two species of the family Syrphidae that are attracted to the flower, mainly Episyrphus balteatus (Shi et al. 2008). The orchid has a yellow spot on the labellum that E. balteatus is attracted to, and after landing on the flower at that location, the fly naturally comes into contact with the pollinia when exiting the flower (Shi et al. 2008). An interesting note about this flower, it was determined by Shi et al. (2008) using

GC/MS to produce no floral volatiles, meaning that visual cues were the main method of attraction over olfactory cues.

In another case, male flies from the family Mycetophilidae visit Pterostylis sanguinea

(Orchidaceae) and attempt to mate with the labellum, as it produces a chemical volatile to attract the males (Phillips et al. 2014). The labellum of this flower is moveable, and is triggered after the fly lands. This motion traps the fly, and as it makes an escape from the flower, it contacts the pollinia and stigma resulting in pollination of the flower (Phillips et al. 2014).

Less common Dipteran pollinators are mosquitoes. Though less frequent as other species, mosquitoes have been documented pollinators of Platanthera obtusata orchids (Thien 1969,

Gorham 1976). Both Aedes communis and Aedes canadensis have been recorded. The pollinia was found to be affixed to the mosquito eyes (Thien 1969).

Lepidoptera

Lepidoptera, the butterflies and moths, are important pollinators for flowering plants. A study by Crepet (1984) categorizes Lepidoptera as the second most important group of pollinators, behind only Apoidea. Even though they are from the same insect order, the pollination syndromes for both butterflies and moths are very different. Because butterflies are active during the day, they are attracted to brightly colored, fragrant flowers. Moths are attracted to fragrance as well, but being night flying insects, they seek out flowers that open in the evening or are more fragrant at night, and are typically white or pale colored (Bawa 1990). One other difference is that butterfly pollinated flowers more often have nectar guides. These are markings on the base of flower petals that act as guides for pollinators, directing them to the nectar source.

Often, these nectar guides are visible only in the ultraviolet spectrum and therefore are invisible to the human eye, but visible to many insects, including butterflies (Thompson et al. 1972).

Both butterflies and moths pollinate a wide range of plant families. Despite being common pollinators, Lepidoptera are less efficient than bees because they have more slender legs, do not have structures for pollen collection, and are not as hairy as many bees are, making it more difficult for pollen to become attached to the insects.

As with many other insect taxa, the relationship between flowering plants and

Lepidoptera can be either generalist or specialist. In some cases, the specialized relationships are very specific, so much that only a single Lepidoptera species can pollinate a flower species.

Darwin's orchid, Angraecum sesquipidale, requires a specific hawk moth (Family: Sphingidae) pollinator to be able to reach into a long nectar spur with its proboscis (Kritsky 1991).

In south Florida, a similarly structured flower, Dendrophylax lindenii, also has a long nectar spur. It is currently hypothesized that another hawk moth species has a similar relationship with D. lindenii due to the nectar spur length (Figure 1-4).

Conversely, other orchids have a generalist relationship with their pollinators, and use other methods to increase the likelihood of successful pollen transfer. There is a hawk moth species in Madagascar (Panogena lingens), that acts as a pollinator for multiple species of angraecoid orchids found there (Nilsson et al. 1987). When specimens of P. lingens were collected, Nilsson et al. (1987) found that pollinia from five orchid species were attached to different areas of the moth. By utilizing different pollinia attachment sites on the pollinator, it would help reduce the accidental placement of pollinia from one orchid species onto the stigma of a different species.

The orchid Epidendrum paniculatum attracts its Lepidopteran pollinator by excreting a chemical attractant. It releases pyrrolizidine alkaloids that male Lepidopterans in the subfamily

Arctiinae, a moth, and subfamilies Ithomiinae and Danainae, butterflies, use for defense and for mate attraction (DeVries and Stiles 1990). Epidendrum paniculatum was surveyed in Costa Rica by DeVries and Stiles (1990), and it was found that 98% of the insects attracted to the flowers were male Lepidopterans that use pyrrolizidine alkaloids.

Fragrances are used for attraction of particular pollinator species. In two very closely related orchids from the genus Gymnadenia, G. conopsea, and G. odoratissima, the fragrance of the flowers was collected in addition to pollinators collected by Huber et al. (2005). They found that there were different physiologically active compounds in each, and that each flower species attracted Lepidopteran pollinators, but that the pollinators of each species did not overlap with each other (Huber et al. 2005).

Sometimes an orchid will attract a pollinator using Batesian mimicry as deceit, such as

Disa ferruginea. The single pollinator of D. ferruginea has been determined by Johnson (1994) to be a Nymphalid butterfly, Meneris tulbaghia. The orchid mimics other flowers, either a red or orange flower based on geographic location of each, that the butterflies commonly visit.

However, unlike the red or orange flowers that they are mimicking, the orchids provide no nectar source for the butterflies (Johnson 1994).

Birds

Though they are the most frequent orchid pollinators, insects are not the only taxon to provide pollination services. Some vertebrates such as birds also may aid in plant pollination, though it is not as common as insect pollination. Avian pollinators require a higher energy output by the plants compared to the quantity of nectar and or pollen required to satisfy insect pollinators (Stiles 1978). Flower-visiting birds are usually less dependent on the plants because they are able to attain their protein and sugar needs elsewhere. In fact, pollination by birds can sometimes be a result of incidental contact to flowers not adapted for avian pollination (van der

Pijl 1961). Typically, bird-pollinated flowers are characterized by their tubular shape and bright, reddish colored flowers, but this is not always an accurate indicator as some bird-pollinated species, especially orchids, have highly different morphologies (Johnson 1996).

Though there are other groups, some of the most common avian pollinators are hummingbirds. Orchids specifically have been documented to have hummingbird-pollinated species. Although only about 3% of orchids are hummingbird pollinated, that still results in the total number of orchid species pollinated by hummingbirds at approximately 1,000 species

(Siegel 2011). Orchids that rely on hummingbirds for pollination are brightly colored and often do not have a fragrance. Many of the orchids pollinated by hummingbirds will produce dark pollinia in shades of gray, blue, or brown. These colors are similar to the colors of

hummingbirds’ beaks, thus reducing the chance of the bird noticing the pollinia and cleaning the pollinia off its beak when grooming (Siegel 2011).

Another example of an orchid-pollinating bird is in the orchid genus Satyrium, with three

South African species confirmed to be bird pollinated. The three species, S. carneum, S. coriifolium, and S. princeps, have many colorful flowers, sometimes up to 100 on each inflorescence, with a large stalk the birds utilize as a perch (Figure 1-5) (Johnson 1996). The pollinators are sunbirds from the genus Nectarinia, small birds with long bills to reach into the flowers for nectar consumption. Though some insects were observed visiting the flowers, they were not capable of removing the pollinia (Johnson 1996). Like with hummingbirds, the orchid pollinia attaches to the bill of the bird as the bird attempts to reach nectar. In most bird-pollinated flowers, a dusting of pollen is left of the birds head. Because of the structure of orchid pollen being compact in a pollinia, it must attach to a birds bill in order for it to be moved to another flower (Johnson 1996).

Other African orchid species from the genus Disa are also sunbird pollinated, but the pollinia is transferred in a different way. Similarly to the Satyrium orchids, D. chrysostachya and

D. satyriopsis flowers, have many flowers along a sturdy inflorescence. When the birds land on the inflorescence, the pollinia is attached to the feet of the birds, and transferred to other flowers as they move between plants (Johnson and Brown 2004). The birds land on the lower flowers and reach up to feed on nectar from the flowers at the top of the inflorescence.

Outside of Africa, an orchid species found on Reunion Island, Angraecum striatum, has been shown to be bird pollinated (Micheneau et al. 2006). Despite A. striatum being self- compatible, it does not self-pollinate and still requires a pollen vector to move pollinia to

produce viable seed capsules. One known vector is Zosterops borbonicus, the Reunion grey white-eye, lands on the flower inflorescence and feeds on nectar (Micheneau et al. 2006).

Bats

Aside from birds, another type of vertebrate pollinator are bats. Bats that pollinate flowers are nectar feeders, and are attracted to large, typically white flowers that are fragrant at night and produce a large amount of nectar (Bawa 1990). Though there are over 500 species of bat-pollinated angiosperms, it is still only about half the number of plant genera compared to bird-pollinated plants (Fleming et al. 2009). Of the 18 families of bats, only two have genera that have evolved morphologically for feeding on nectar, Phyllostomidae and Pteropodidae (Fleming et al. 2009). Despite insects being the leading biotic pollinators, there are benefits to having larger pollinators such as bats. Bats are much larger, meaning they can carry larger amounts of pollen, and they fly much greater distances than an insect would (Bawa 1990, Fleming et al.

2009). While some bats are destructive eaters, destroying the flowers in the process of pollination, others simply hover near the flower and drink nectar (van der Pijl 1961).

One plant family that benefits from bat-pollination is the Bromeliaceae. Bromeliads are uncommon as a plant family in their pollination because vertebrates are more common pollinators than insects (Sazima et al. 1989). Encholirium glaziovii is a bromeliad found in Brazil that attracts the nectar-feeding bat Lonchophylla bokermanni (Sazima et al. 1989). The bat feeds while hovering, and as a result, the snout of the bat becomes dusted with pollen.

The passion flower Passiflora mucronata, also in Brazil, is another example of a bat- pollinated flower. Other visitors to the flowers include Lepidoptera, Hymenoptera, and some hummingbirds, but only two species of bats, Glossophaga soricina and Carollia perspicillata, were effective pollinators (Sazima and Sazima 1978). Some other plant families that have bat- pollinated species include Bombaceae, Mimosaceae, and Caesalpiniaceae (Bawa 1990). These

bat-pollinated flowers also usually have a higher amount of nectar produced than insect- pollinated flowers (Bawa 1990).

Orchids and Orchid Pollination

The Orchidaceae is one of the largest families of flowering plants in the world, with almost 30,000 species (Brown 2005). Specialized selection pressure on some orchid taxa has resulted in evolution of dramatic morphological differences between groups (Kew Royal Botanic

Gardens 2015). One trait that has evolved in orchids is the fusion of the stamens with the pistil in the flowers. Reduction in stamen number has led to orchid groups with three, two, or one stamen.

Over 99% of all species of orchids only have one stamen in the flower, which is one of the major features of the Orchidaceae (Kew Royal Botanic Gardens 2015). Orchids have the same basic reproductive structures across most species (Figure 1-6). There is a central structure referred to as the column, which houses both the male part of the flower, the anther, and the female part of the flower, the stigma (Roberts and Dixon 2008). Below the column is a petal that has been modified into a labellum or lip, which acts as a landing area for pollinators and directs them to the nectar source within the flower (Brown 2005). As a pollinator moves towards the nectar source, it comes into contact with the pollen. Unlike the loose pollen grains of most flowering plant families, orchid pollen is housed in compact structures called pollinia. The pollinia are located in the male anther cap, and there is often a sticky structure called the viscidium that helps the pollinia adhere to a pollinator as it nectars (Roberts and Dixon 2008). The stigma is located just on the underside of the column. When a pollinator visits another flower, the pollinia will be transferred to the stigma. If pollination is successful, the ovary of the flower will begin to swell and form a seed pod filled with millions of seeds (Roberts and Dixon 2008).

Because orchids are so diverse, the pollination systems are highly variable and sometimes extremely specialized. Many orchids use some form of deception to attract pollinators, often by either physical appearance or chemical cues, as discussed in the previous section (Wong and

Schiestl 2002, Jersáková et al. 2006). The most common example of deception using the physical appearance of the flowers is by Ophrys apifera, or the bee orchid. The lip of this flower resembles a female bee sitting on the flower. Male bees are deceived into attempting to copulate with the flower, and eventually move on in search of other females. Through these mating attempts, the pollinia attaches to the male bee. When the male is deceived by another flower, the pollinia transfers to the pollen receptor, resulting in pollination (Devey et al. 2008). In addition to using visual tricks, many orchids release compounds that attract pollinators. The compounds may resemble the pheromones of a mate, or release a chemical that the pollinator can use for defense or in mate attraction (Wong and Schiestl 2002). One example is the orchid Epidendrum paniculatum. It releases pyrrolizidine alkaloids that male Lepidoptera in the family Arctiidae and

Nymphalidae subfamilies Ithomiinae and Danainae use for both seeking mates and defense

(DeVries and Stiles 1990). DeVries and Stiles (1990) surveyed E. paniculatum in Costa Rica along with pyrrolizidine alkaloid baits and found that 98% of the insects attracted to the flowers and baits were male Lepidoptera that use pyrrolizidine alkaloids for attracting mates and for predator defense.

Orchids have a variety of relationships with pollinators. Some species are pollinated by a diverse set of insects and other taxa, while others are specialized with only a single pollinator species. The advantage for orchids that have specialized relationships with a single species, is that it often results in more efficient pollen transfer than those species with multiple general pollinators. Specialization causes more direct transfer from one flower to another, while reducing

the chance that the pollinia will be dropped or transferred to the wrong species (Scopece et al.

2010). A species of orchid known as Angraecum sesquipedale, Darwin's orchid, and an orchid native to south Florida, Dendrophylax lindenii, the ghost orchid, both require a particular lepidopteran pollinator. Hawkmoths have a long proboscis, and these orchids have a long nectar spur that the hawkmoth is able to reach (Kritsky 1991). Orchids may also attach pollen to a specific location on the pollinator's body, ensuring that even if the pollinator visits another species, the pollinia likely will not be transferred until it revisits an orchid of the same species

(Scopece et al. 2010).

Flowers of the Orchidaceae family are known to be pollinated by a diverse set of pollinator taxa. Bees and wasps in the order Hymenoptera are the most common orchid pollinators, but Lepidoptera, Diptera, Coleoptera, and other insect orders are known orchid pollinators (Statman-Weil 2001, Lehnebach and Robertson 2004, Micheneau et al. 2010, Stökl et al. 2011). Hymenoptera are the most diverse insect order of pollinators. In this order, the euglossine bees (the orchid bees) have one of the closest relationships with orchids. The orchid releases a compound that attracts male bees. The compound does not mimic mating pheromones, but the bee collects the compound and then uses it to attract females (Williams and Whitten

1983, Ackerman 1983). The bee often becomes temporarily trapped inside the flower, where the orchid's pollinia attach to the bee. Then, when the bee visits another flower to gather the compounds, the pollinia is transferred and the orchid is fertilized (Williams and Dodson 1972,

Ackerman 1983, Ramirez et al. 2010).

Though they are the most common, insects are not the only type of orchid pollinators.

Hummingbirds are the primary avian pollinators of orchids. Although only about 3% of orchid species are hummingbird pollinated, this still represents almost 1,000 known species (Siegel

2011). Orchids that rely on hummingbirds for pollination are brightly colored and often do not have a fragrance. Many of the orchids pollinated by hummingbirds will produce pollinia that are darker in color and nondescript, so that when it attached to the bill the bird is less likely to remove it by cleaning the beak when it sees a sharp color contrast (Siegel 2011). A recent study found that a genus of hummingbird pollinated orchids had significantly lower fruit set in natural conditions than when hand-pollinated (Nunes et al. 2015).

There are over 200 species of orchids in North America and of these, 106 are found in

Florida (Brown 2005). Of these 106 species, over half are restricted to the southernmost areas of

Florida, mainly the Big Cypress Basin (Zettler et al. 2012). About half of the orchid species in

Florida are listed as either state endangered (56) or threatened (17), with several more being proposed to be added to these lists (Brown 2005). Numerous protected areas in south Florida

(e.g., Everglades National Park, Fakahatchee Strand, Florida Panther National Wildlife Refuge, and Big Cypress National Preserve) provide habitat for these state endangered orchids. Despite this, the threats of habitat degradation, invasive species, and poaching are still present. Several species of threatened and endangered species thrive in these protected areas such as Epidendrum nocturnum (night-fragrant epidendrum), Epidendrum rigidum (rigid epidendrum), Encyclia tampensis (Florida butterfly orchid), Epidendrum amphistomum (dingy-flowered star orchid),

Prosthechea cochleata (clamshell orchid), Dendrophylax lindenii (ghost orchid), Polystachya concreta (helmet orchid), Campylocentrum pachyrrhizum (cigar orchid), and Harrisella porrecta

(leafless harrisella) (Stewart and Richardson 2008, Ray et al. 2012). It is important to understand the pollination system of orchids in order to develop plans for conservation for threatened and endangered species.

In spontaneous autogamous systems (auto-pollination), the pollinia are located on the stigma and pollination occurs in the absence of a pollinator. This process occurs through several mechanisms including powdery pollinia, expansion of the pollinia, or a deformation of part of the column (Catling 1990, Gale 2007).

Through autogamy, orchid species that exhibit this trait are unrestrained by limitation imposed by the need to co-occur with a suitable pollinator. It is thought that autogamy facilitates range expansion of a plant into areas where pollinators are rare or not present, and there is a possibility autogamy is the consequence of range expansion (Catling 1990). Fenster and Martén-

Rodriguez (2007) suggest that autogamy could be more widespread than previously believed, and auto-pollination could have evolved alongside self-pollination to adapt to selection pressures. These pressures include pollinator limitation and the ambiguity of seed set, as auto- pollination can provide reproductive insurance against the specialized pollinators that many orchids have (Fenster and Martén-Rodriguez 2007).

Autogamy is thought to be somewhat common throughout the Orchidaceae (Johnson et al. 2009). There are more than 350 orchid species that have been reported to exhibit autogamy.

For example, it is estimated that at least 20% of species found in Ecuador and Puerto Rico are autogamous (Catling 1990). Several species of orchids found in southern Florida have been observed to exhibit autogamy. In these reports, seed capsule formation and see viability was not investigated, making it difficult to determine if autogamy is an advantageous method of seed capsule production (Goss 1973, Catling 1987, Johnson et al. 2009). A study by Johnson et al.

(2009) examined the breeding system of a Florida terrestrial orchid Eulophia alta, which has been reported as autogamous. In those occurrences of auto-pollination, they found that only 7.1% of the flowers formed seed capsules, and this data indicated that autogamy is rare in E. alta

(Johnson et al. 2009). This study suggests that the capsules formed naturally in the field may result from pollination events that were not observed.

The goals of this research were to determine the floral fragrance of P. cochleata, identify potential pollinators of both P. cochleata and E. tampensis, and determine if these species are capable of self-pollinating, and to compare the percent germination of seeds from resulting seed capsules of both P. cochleata and E. tampensis. Because of the protected status of these flowers, it is important to learn about their pollination and reproductive biology and use that information to improve future conservation efforts for these orchids. It's unknown if either of these species could have future benefit to man, studies into their potential for medical purposes have yet to be conducted.

Figure 1-1. The basic structure of a flower including both male and female reproductive parts. Illustration by Fauve L. Wilson.

Figure 1-2. Map of Florida representing conservation land and strategic habitat necessary for maintaining long-term biodiversity (Kautz and Cox 2001).

Figure 1-3. Diagram highlighting the pollen basket on the leg of a honey bee (Apis mellifera) (HymAToL 2006).

Figure 1-4. Dendrophylax lindenii, the ghost orchid, in south Florida. The red arrow is pointing to the flowers nectar spur, which suggests a hawk moth pollinator with a long proboscis. Photograph by Haleigh A. Ray.

Figure 1-5. Inflorescences of Satyrium coriifolium orchids in bloom. Photograph: Wikimedia commons.

Figure 1-6. Structure of a flower in the family Orchidaceae (Subrahmanyam 2010).

CHAPTER 2 FLORAL FRAGRANCE ANALYSIS OF Prosthechea cochleata (ORCHIDACEAE), AN ENDANGERED NATIVE, EPIPHYTIC ORCHID, IN FLORIDA

Introduction

The fragrance produced by flowers is one of the primary attractants of pollinators to a flower. Orchids specifically have a diverse range of pollination systems, including approximately

10,000 species that use deception to attract pollinators (Dudareva and Pichersky 2006). Many orchid species that are bee-pollinated use fragrance as a dominant attraction method, along with flower color and structure as reinforcement of the signal (Dodson et al. 1969). A flower's fragrance is often a combination of compounds, which can belong to numerous chemical classes

(Knudsen et al. 1993). Typically, pollination by insects requires floral fragrances for pollinator attraction (Altenburger and Matile 1988). Studying the fragrance produced by flowers is an important component in understanding plant-pollinator interactions.

Floral fragrance analysis of Dendrophylax lindenii, the ghost orchid, an epiphytic orchid found in south Florida, has shown that (E)-β-ocimene is one of the most abundant volatile organic compound produced by this orchid, as well as the presence of α-pinene (Sadler et al.

2011). Three D. lindenii flowers were sampled, each with similar results. A study of another orchid, Epidendrum ciliare, found high fragrance variation between plants of the same species

(Moya and Ackerman 1993).

The purpose of this study was to identify the fragrance compounds from the clamshell orchid, Prosthechea cochleata (Figure 2-1). In the United States, this state endangered orchid is found growing epiphytically in southern Florida. According to Cancino and Damon (2006), flowers of P. cochleata found in Mexico did not produce any volatiles when flowers were sampled with the protocol developed by Heath and Manukian (1992). My Florida data collected

from using the same sampling procedure indicated that P. cochleata does produce volatiles, including α-pinene, (E)-β-ocimene, benzaldehyde, mesitylene, and nonanal.

Materials and Methods

Fragrance Collection

Volatiles were collected using a procedure originally described by Heath and Manukian

(1992), and modified by Cancino and Damon (2007) and Sadler et al. (2011). Using this method, samples could be collected repeatedly from the same flower over several days, without removing the flower from the plant. A Reynolds® oven bag was used to cover the flower, and sealed around the inflorescence to prevent ambient air from entering the bag during sampling (Figure 2-

2). The oven bags were used once, and then decontaminated by washing with 75% ethanol and baking in an oven at approximately 93ºC for 30 minutes. This prevented any residual volatiles from a sample being detected in a subsequent sample. For each sample, volatiles were collected for 10 minutes. Dry charcoal filtered air was pushed into one end the oven bag and exited the chamber via a vacuum system. The air then passed through a volatile collection filter containing

50 mg of Tenax® Porous Polymer Adsorbent (Sigma-Aldrich, USA).

Four individual plants were sampled by collecting volatiles from one flower per plant.

The sampled plants included three growing in pots in a greenhouse at the University of Florida

Entomology and Nematology Department (UF) in Alachua County, Florida, and one wild orchid growing in situ at the Florida Panther National Wildlife Refuge (FPNWR) in Collier County,

Florida. The orchid in the FPNWR was found growing epiphytically, approximately 1.75 m above the ground. The collection equipment was elevated to accommodate this height. The beginning of flower sampling started on the first day that a new bloom was open, which was deemed Day 0.

The flowers from the UF greenhouse were sampled on days 0, 5, 10, and 15. Flowers were sampled twice, at 8:00 AM and 4:00 PM, on days 0 and 10; and three times, at 8:00 AM,

4:00 PM, and the following 12:00 AM EST, on days 5 and 15. Volatile collection occurred within ±15 minutes of the scheduled time. Sampling the flowers at different time points allowed us to detect any fluctuation in fragrance strength throughout the day. This resulted in 10 total floral samples for each of the three UF flowers, for a total of 30 floral samples from the greenhouse flowers.

The in situ P. cochleata orchid at FPNWR was sampled on two different days. The flower selected was determined to have opened most recently, by observing the flowers and noting when a new bloom had opened. The sampling schedule on day 0 and day 5 was the same as sample collection from greenhouse flowers at UF, for a total of five floral samples collected of the FPNWR flower. This method yielded 35 total floral samples collected from the three UF flowers and the one FPNWR flower (Table 2-1).

Three control samples (plant, substrate, air) were collected. One by sampling the plant excluding all flowers (leaves, stems, pseudobulbs), a second by sampling either the plastic container and growing medium used for the greenhouse plants or part of the tree for the epiphytic orchid in FPNWR, and a third by sampling the ambient air in the place of collection. Peak volatiles collected from control samples were compared with the peak volatiles from the floral samples to identify and remove any peaks that do not originate from the flower. Ambient air control samples were collected at each sampling date (N=14), and plant and substrate samples were collected at 8:00 AM EST on the second sampling day for each plant (N=4 for both plant and substrate samples). This resulted in r a total of 22 control samples from the three UF flowers and the one FPNWR flower (Table 2-1).

Identification of Floral Odor

After the floral and control samples were collected, they were labeled and sealed using

Teflon thread seal tape and stored in a freezer (-80 ºC) until they were analyzed. The volatile compounds collected were analyzed at the United States Department of Agriculture (USDA) in

Gainesville, Florida by gas chromatography mass spectrometry (GC/MS) [GC: Agilent 6890 with an HP-5MS capillary column of 30 m long, 0.25 mm inner diameter, and 0.25-µm film thickness; MS: Agilent 5973 mass selective detector, 70 eV, equipped with a thermal desorption cold trap injector (TCT) (CP4010; Chrompack, Bergen op Zoom, The Netherlands)]. Headspace volatiles collected on Tenax ® TA were released from the adsorbent by heating in the TCT at

220°C for 8 minutes within a flow of helium gas. The desorbed compounds were collected in the

TCT cold trap unit (SIL5CB-coated fused silica capillary) at –130°C. Flash heating of the cold trap unit injected the compounds into the capillary column of the gas chromatograph to which the cold trap unit was connected.

The oven temperature of the GC was programmed to rise from 40°C (5-min hold) to

280°C at 15°C/min. The headspace volatiles were identified by comparing their mass spectra to those of the database (Wiley7N and Wiley275) and by comparing their retention times to those of authentic compounds. Volatiles were identified by comparison of mass spectra (a) with mass spectra libraries (NIST and Department of Chemical Ecology, Göteborg University, Sweden) and (b) with mass spectra and retention times of authentic standards. After the results were obtained, the volatile peaks were compared to each other to locate the common peaks in the flower samples. These volatiles were then referenced on Pherobase, an online database of pheromones and semiochemicals.

Results

Analysis of the fragrance by GC/MS indicated several compounds repeatedly appearing in the floral volatiles of P. cochleata (Table 2-2). The samples from all four flowers presented similar volatile analyses. All of the chemical compounds listed were compared to those on

Pherobase and prior floral fragrance publications, with many found in floral fragrances from other plants in the Orchidaceae family (Knudsen et al. 2006). The most abundant volatiles collected were Pseudocumene, Nonanal, and Mesitylene, respectively.

Variation in fragrance samples from 8:00 AM, 4:00 PM, and 12:00 AM were noted.

According to the GC/MS analysis, the abundance was lowest during the 12:00 AM collection period, while the 8:00 AM and 4:00 PM samples resulted in numerically higher abundances. This trend was also noted by human olfaction in the laboratory at the same time periods. There were also slight differences in the amount of certain volatiles between the three plants, most noticeably in the Pseudocumene.

Discussion

The results of this study contrast those of Cancino and Damon (2006), by showing several floral fragrance compounds detected in P. cochleata. A recent study on the floral volatiles of a terrestrial orchid, Gymnadenia odoratissima, determined that the floral compounds of these flowers differed by location, being stronger in lowlands than in the mountains (Gross et al. 2016). The region of the flower could be influencing the volatiles produced, as G. odoratissima produces differing compounds based on the location of the plants.

The differences in volatile percentage over the duration of the study, such as with

Pseudocumene, could be explained by the overall fragrance changing over time. In a study by

Ackerman (1989), it was found that several species of euglossine bees, also known as orchid bees, exhibited a fragrance preference between varying seasons. The collections were done as the

flowers from each plant opened, which took place over the course of 8 to 10 weeks. As for the lower fragrance emission at night (12:00 AM) compared to 8:00 AM and 4:00 PM, this suggests that a pollinator would be active during the day, and that the flower would not be as attractive to pollinators active in the evening. Pollination requires the floral fragrance emissions to be synchronized with the activity of insect pollinators, so that successful pollination will be maximized (Altenburger and Matile 1988).

An interesting aspect of the reproductive biology of P. cochleata is that this flower is thought to be autogamous, or self-pollinating (Higgins 2003). It takes energy for a flower to produce a fragrance, which seems counterproductive for a flower species that is autogamous to be producing fragrance volatiles (Dudareva and Pichersky 2006). Additionally, the fragrance declined during the evening hours, suggesting that it would be most attractive to a day-flying pollinator.

In a review of floral scents published in 2006, there were an estimated 417 species in the

Orchidaceae from which fragrance had been analyzed (Knudsen et al. 2006). Although this was a significantly higher number than the next family (Araceae- 55), it is still a small percentage of orchid species worldwide, as there are nearly 30,000 known orchids (Brown 2005). The review by Knudsen et al. (2006) also noted the top five fragrance compounds found in plant species (α- pinene, (E)-β-ocimene, limonene, linalool, and myrcene), the first three of which were identified in P. cochleata. It is important to understand the fragrance aspect of P. cochleata reproductive biology, as it could lead to better conservation efforts for this state endangered orchid. By knowing the composition of the floral fragrance, it could allow for better methods of pollinator attraction such as augmenting fragrance by introducing a blend of the compounds found being

produced by the flowers. An increase in the pollinators would result in more seed capsule production, and elevate the number of flowering plants in a location.

Table 2-1. Sample schedule for each of three Prosthechea cochleata flowers from different plants at the Entomology and Nematology Department (UF), and the single flower at the Florida Panther National Wildlife Refuge (FPNWR). Additional samples for three controls were collected, the plant, substrate, and the air control (Control- Air*a, Control- Plant *p, Control- Substrate*s). Day 0 Day 5 Day 10 Day 15 Total Samples *a *a,*p,*s *a *a UF 1 8:00 AM 8:00 AM 8:00 AM 8:00 AM 16 4:00 PM 4:00 PM 4:00 PM 4:00 PM 12:00 (10 floral) 12:00 AM AM *a *a,*p,*s *a *a UF 2 8:00 AM 8:00 AM 8:00 AM 8:00 AM 4:00 16 4:00 PM 4:00 PM 4:00 PM PM 12:00 AM (10 floral) 12:00 AM *a *a,*p,*s *a *a UF 3 8:00 AM 8:00 AM 8:00 AM 8:00 AM 16 4:00 PM 4:00 PM 4:00 PM 4:00 PM 12:00 (10 floral) 12:00 AM AM *a *a,*p,*s FPNWR 8:00 AM 8:00 AM ______9 4:00 PM 4:00 PM (5 floral) 12:00 AM

Table 2-2. A list of the top eight compounds detected in the floral fragrance samples of Prosthechea cochleata. Samples include three flowers from the University of Florida greenhouse (UF 1-3), the flower from a wild plant at the Florida Panther National Wildlife Refuge (FPNWR), and an air control. Air Compound UF 1 UF 2 UF 3 FPNWR Sample Benzaldehyde  (2.48)  (2.64)  (1.54) -- T Mesitylene  (4.04)  (1.41)  (1.86)  (1.75) T α-pinene  (1.94)  (2.20)  (1.62)  (1.80) -- (E)-β-ocimene  (1.80)  (1.42)  (1.54) -- -- Nonanal  (4.13)  (2.31)  (3.88)  (2.7) -- Decanal  (2.14)  (1.53)  (2.54) -- -- Pseudocumene  (5.17)  (3.57)  (11.55)  (2.69) T Limonene  (2.15) T  (1.01) -- -- *Number listed in parentheses after the  refers to the average abundance of the corresponding volatile across all floral samples for that plant *T represents only trace abundance (< 1%) *-- refers to the corresponding compound not being found in that sample

Figure 2-1. Flower of Prosthechea cochleata, the clamshell orchid, growing at the Florida Panther National Wildlife Refuge. Photograph by Larry W. Richardson.

Figure 2-2. Image showing the collection of floral volatiles from the greenhouse grown Prosthechea cochleata. Photograph by Haleigh A. Ray.

CHAPTER 3 POLLINATION OF Encyclia tampensis, THE COMMERCIALLY EXPLOITED BUTTERFLY ORCHID, BY VARIOUS INSECT TAXA IN SOUTH FLORIDA

Introduction

Members of the flower family Orchidaceae, the orchids, are known to be pollinated by a diverse set of taxa. The most common orchid pollinators are bees and wasps in the order

Hymenoptera. However, insects in Lepidoptera, Diptera, Coleoptera, and other orders are known orchid pollinators as well (Statman-Weil 2001, Lehnebach and Robertson 2004, Micheneau et al.

2010, Stökl et al. 2011). Orchids have various relationships with pollinators. Some species are pollinated by multiple pollinator species, while others are pollinated by only a single species.

Orchid flowers have the same basic floral structures for pollination across most species. There is a central structure referred to as the column and it contains both the male part of the flower

(anther) and the female part of the flower (stigma) (Roberts and Dixon 2008). Below the column, a petal that has been modified into a labellum or lip that acts as a landing area for pollinators.

This directs them to the nectar source within the flower, causing them to contact the orchid pollen (Brown 2005). Unlike loose pollen grains produced by most flowering plant families, orchid pollen is housed in compact structures called pollinia. Pollen transfer occurs when the pollinia is attached to a visiting insect and transferred between flowers. When pollination occurs, the ovary will begin to swell and form a seed capsule filled with millions of seeds (Roberts and

Dixon 2008).

Florida is home to over 100 species of orchids, typically in the southernmost areas, as these locations provide ideal growing conditions for many epiphytic species. Several protected parks and refuges in south Florida provide habitat for these species, over half of which are listed as threatened or state endangered (Stewart and Richardson 2008). Despite these protected areas, many orchids still face threats from habitat degradation, invasive species competition, poaching

of plants, and pests. In a survey of orchid species in south Florida, E. tampensis was found to have Boisduval scale (Diaspis boisduvalii) present on some of the adult plants (Ray et al. 2012,

Zettler et al. 2012). Currently, there is little information regarding the pollination of the Florida butterfly orchid, Encyclia tampensis (Figure 3-1), which is listed as commercially exploited. A better understanding of this orchid’s pollinator(s) will facilitate science-based future conservation decisions.

Observations of E. tampensis at the Florida Panther National Wildlife Refuge (FPNWR) in Collier Co., FL have shown a higher seed capsule formation of flowers near the refuge work center compared to a lower seed capsule formation of flowers found near secluded ponds. The orchids found growing epiphytically in trees in the ponds are surrounded by fewer floral resources than those near the work center, where there are a variety of flowering plants. The objective of this study was to identify the potential pollinator(s) of E. tampensis, compare seed capsule formation between flower locations, and identify factors that influence differences in seed capsule formation.

Materials and Methods

Pollinator Exclusion

This study was conducted at the Florida Panther National Wildlife Refuge (FPNWR) in

Collier County, Florida (26.171577, -81.347108), during three blooming seasons (2015-2017).

Four separate locations at the refuge were selected (Figure 3-2). Each location had three geographically distinct sites, spaced at least 10 m apart, where E. tampensis orchids occur naturally. The GPS coordinates of these locations have been withheld due to the threatened status of these orchids and others growing in the area. Location 1 was a developed, landscaped work center at FPNWR, while Locations 2-4 were all natural freshwater wetland forest habitat. Each site had at least three blooming E. tampensis, with a total of five or more flowers per site. The

number of flowers blooming at each site was recorded. To determine if the orchid requires a pollinator for seed capsule production, mesh exclusion bags were placed over at least three unopened or newly opened, unpollinated flowers at each location (Figure 3-3). When exclusion bags were placed over a flower, they were typically covering an inflorescence of multiple flowers, not a single flower per bag. Evidence of previous pollination was determined by visually inspecting the flower stem. There is a noticeable difference in pollinated flowers, as seed capsule production begins quickly after fertilization (Figure 3-4). The flower stems change color from a yellow flower color when unfertilized to a dark green, and begin to swell as the seed capsule develops.

Pollinator Collection

During the study, potential pollinators were collected using traps as well as active sampling. Three types of traps were used to collect insects: Blue vane traps (SpringStar ®,

Seattle, WA), WHY (wasp, hornet, yellow jacket) traps (Rescue ®, Spokane, WA), and colored insect bowl traps comprised of three bowls (Blue Sky®, Brooklyn, NY): blue, yellow, and white

(Figure 3-5). Three traps of each type were set up at each location within 5 m of flowering E. tampensis orchids. The blue vane and WHY traps were suspended from branches at approximately 1.5 m above the ground, while the bowl traps were placed at ground level.

Though the orchids are epiphytic and can grow lower or much higher, 1.5 m was chosen to allow traps to easily be hung and checked. Each trap contained approximately 200 ml of water with

0.01% of the surfactant Silwet L-77 (Helena ®, Collierville, TN) to break the surface tension and prevent insects from escaping. Traps were checked daily and insects were collected over a six- day period each year during peak blooming between 1300 and 1600 h. The traps were available to insects all hours of the day.

In addition to trapping, all locations were actively sampled twice each day in 45-minute increments (15 minutes at each site), for a total of 90 minutes of observation at each location per day. When actively collecting insects, the locations were visited in a varying order so that they were being monitored at various times throughout morning or afternoon (Example: Day 1-

Location 1, 2, 3, 4; Day 2- Location 2, 3, 4, 1; etc.). This was to prevent visiting one specific location earliest in the morning or close to the afternoon during every collection period. Insects that landed on the floral blooms were collected using an aerial net. Collected specimens were identified and released if possible, or preserved by freezing and taken to the Entomology and

Nematology Department at the University of Florida (UF/IFAS laboratories) for identification.

All flowering plants within a 10 m radius from the orchids were photographed and identified.

This study was repeated for three years when the flowers were blooming, mid-May to early June, from 2015-2017.

Statistical Methods

I used an analysis of variance (ANOVA) to compare any significant differences between pollinator species at each collection site at FPNWR and differences in seed capsule formation. A t-test was performed to test for any difference in pollinator abundance in the morning collections compared to the afternoon collections. These statistical tests were completed using JMP

Statistical Analysis Software. For complete data sets see Appendix A.

Results

Pollinator Exclusion

During the three year study period, the average total numbers of flowers studied ± SD at each location were 125 ± 35, 97 ± 17, 50 ± 3, and 53 ± 5 at Locations 1, 2, 3, and 4, respectively.

Each location had a total of three sites within, for a total of N = 9 at each location (N = 36 total) over the course of the study. During the three years of the study, a total of 231 flowers (about

24% of all flowers) were covered with mesh exclusion bags across all locations at FPNWR. Of these, zero seed capsules formed from bagged flowers, making it unlikely that these flowers are capable of self-pollination. Of the flowers that were left uncovered, the flowers at the developed location (Location 1) produced a significantly higher percentage of seed capsules (P < 0.0001)

(Table 3-1).

Pollinator Collection

Over the three-year collection period, a total of 46 insects were captured by active sampling, and 83 were captured in the three different traps. Mosquitoes or non-insect arthropods collected in the traps were excluded from the study due to the unlikelihood that they would be pollinators of this species. However, specimens were still checked for visible signs of pollinia before being discarded (Figure 3-1), as mosquitoes have been recorded as pollinators of other orchid species (Statman-Weil 2001). The insects collected by active sampling consisted of three orders, Hymenoptera, Diptera, and Coleoptera, with Hymenoptera being the principal order collected. This was true for each of the four locations, with Hymenoptera being most common, followed by Diptera and Coleoptera respectively. Insects that were repeatedly net collected over the course of the study were Trigonopeltastes delta (Coleoptera), Copestylum sexmaculatum

(Diptera), and several Bombus spp. (Hymenoptera). There were five orders present in the different trap types, Hymenoptera, Diptera, Coleoptera, Lepidoptera, and Hemiptera. Because active sampling occurred by net collecting insects that were visiting the flowers, those are much more likely to be representative of the actual pollinators of E. tampensis.

The higher seed capsule formation typically seen at Location 1 could be influenced by a higher number of pollinators present at that location. The number of insects actively collected at

Location 1 was significantly higher than that at any other location over the three-year period (P =

0.0057) (Figure 3-6). However, there was no significant different in the quantity of insects

collected in each of the three orders (Figure 3-7). Additionally, there were more blooming flowers at Location 1 compared to any other location. While Locations 2-4 consisted of swamps full of pop ash (Fraxinus caroliniana), pond apple (Annona glabra), and cypress trees

(Taxodium distichum) with no other blooming flowers besides the orchids, Location 1 had several species in bloom (Table 3-2).

Insect traps were checked once daily in the afternoons, but any insects that were collected by active sampling were sorted into those collected in the morning (0800-1200) or those from the afternoon (1230-1630). Overall, there were 24 insects captured in the morning and 22 insects captured in the afternoon over the three-year study (Figure 3-8). At Location 1, there were almost twice as many insects collected in the morning (19) than in the afternoon (10). Three, two, and zero insects were collected from Locations 2, 3, and 4 in the morning, and 6, 3, and 3 were collected in the afternoon, respectively. Based on the results of a two-sample t-test, there was no significant difference between the number of insects caught in the morning and the number caught in the afternoon (t=0.877).

In addition to the 46 insects actively collected, there were 83 more collected from either the blue vane, W.H.Y., or colored bowl traps. Of those 83, only 17 of them were species also collected during active sampling, leaving 66 specimens specific to the traps. Across all four locations, Hymenoptera and Diptera were the most prominent orders found in these traps (P =

0.02), with no significant difference in abundance between the two (P = 0.46). When comparing the orders collected in various traps, Diptera were only collected in the colored bowl traps, mostly yellow and white, while Hymenoptera were collected in all but blue pans. Though found in all traps, Hymenoptera were collected in highest numbers in the blue vane traps.

Discussion

I conducted this study to begin to understand the pollination biology of E. tampensis orchids in Florida. Though restoration efforts are beginning, threats of habitat loss, pests, and poaching are increasing for this orchid. My data suggest that flowers of E. tampensis are generalist pollinated, likely by multiple insect orders, as the three most common genera actively collected from flowers were Trigonopeltastes (Coleoptera), Copestylum (Diptera), and Bombus

(Hymenoptera). However, future work could be done to verify this if pollinators could be collected with E. tampensis pollinia attached to their body. The delta flower scarab, T. delta, has been recorded as a flower-visiting species across Florida and in other studies as well, including records of activity in Everglades National Park, located near my field site (FPNWR) (Fontes et al. 1994, Pascarella et al. 2001). The six-spotted bromeliad fly, Copestylum sexmaculatum, was recorded in south Florida from the same survey of Everglades National Park, and in a survey of

Archbold Biological Station, both categorizing it as a flower-visiting species (Pascarella et al.

2001, Deyrup and Deyrup 2012). Bombus spp. (bumble bees) are known generalist bee pollinators.

While the same three genera were collected throughout the refuge, they, along with all other actively collected flower visitors, were most abundant at Location 1 at FPNWR. As noted,

Location 1 was the developed work center and represented a disturbed habitat. Shown in Table

3-2, there were several species of blooming plants at Location 1 that were not present in

Locations 2-4, which may have been a factor in the increased insect activity. Additionally,

Location 1 had a significantly higher number of flowers resulting in seed capsules during two of the three years of the study. It is possible that planting other native, flowering plants near the edges of the swamp habitat could increase pollinator activity in those locations.

In addition to identifying the potential pollinators of E. tampensis, it was also determined that this species is not capable of self-pollination. Other orchid species, such as Epidendrum nocturnum, the night-fragrant orchid, are capable of either autogamy, self-pollination, or cleistogamy, self-pollination without the flower opening first (Stort and dos Santos Pavanelli

1985, Brown 2005). Self-pollination can be advantageous for plants with a short flowering period, limited presence of pollinators, or competition for pollinators (Wyatt 1986, Snell and

Aarssen 2005). A disadvantage is that self-pollination could increase the rate of inbreeding in plants, and reduce the fitness of the population (Jersáková and Johnson 2006). It is possible that there is no advantage for the evolution of self-pollination in this species, because E. tampensis is likely pollinated by a broad range of insects. After self-pollination experiments were performed on Disa pulchra, an orchid that is only fly pollinated, it was found that the resulting seed capsules had about half the number of viable seeds compared to seed capsules formed from cross-pollinated plants (Jersáková and Johnson 2006). Disa pulchra does not provide a nectar reward for its fly pollinators, possibly causing flies to visit fewer flowers on the same plant and decreasing the likelihood of self-pollination.

Our study shows that while E. tampensis is not capable of self-pollination, it is potentially pollinated by a range of flower-visiting insects. Furthermore, my data suggest that by having a variety of flowering plants nearby, pollinator activity on E. tampensis may be increased. Not only were more pollinators collected at the developed location of FPNWR, but this location also produced more seed capsules from the open flowers. Further studies are underway to examine the viability of seeds from the E. tampensis seed capsules collected during this research. Overall, this research provides a better understanding of the reproductive requirements of E. tampensis flowers in south Florida. Future research may be directed towards DNA analysis of gut contents

of the suspected pollinators, searching for the presence of E. tampensis pollen. Having this information will be useful for conservation efforts for these orchids, both for protecting current populations and establishing new populations in south Florida.

Table 3-1. Three year mean percentage of seed capsule formation (±SD) for flowers at each of the locations where Encyclia tampensis was studied at the Florida Panther National Wildlife Refuge. Location 1 was at a developed work center, while the other locations were in the natural refuge habitat. Location 1 had the highest percentage of seed capsule development each year. Row means with the same letter are not different at α ≤ 0.05. Seed Capsule Formation Location 1 Location 2 Location 3 Location 4

2015 25.6% ± 1.8a 10.3% ± 2.3b 8.8% ± 3.5b 3.2% ± 2.9b

2016 28.5% ± 7.3a 14.3% ± 2.2b 9.3% ± 8.5b 8.2% ± 2.6b

2017 21.2% ± 9.5b 16.8% ± 1.5b 5.8% ± 5b 5.5% ± 0.8b

Table 3-2. Identified plants that were in bloom at each of the four locations across the Florida Panther National Wildlife Refuge that were within 10 meters of blooming Encyclia tampensis flowers. Location 1 was a developed, landscaped work center in the refuge, while Locations 2-4 were natural swamp habitat. Location 1 Location 2 Location 3 Location 4

Fraxinus caroliniana (Pop Ash) X X X X

Annona glabra (Pond Apple) X X X

Taxodium distichum (Pond Cypress) X X X

Campis radicans (Trumpet Vine) X

Seville orange (Bitter Orange Tree) X

Bidens alba (Spanish Needle) X

Allamanda cathartica (Allamanda Vine) X Heliconia latispatha (Expanded X Lobsterclaw Heliconia) Catharanthus rosea (Rosy Periwinkle) X

Figure 3-1. Butterfly orchid (Encyclia tampensis) and orchid pollinia. A) Encyclia tampensis, the Florida butterfly orchid, growing at the Florida Panther National Wildlife Refuge in Collier Co., Florida. Photograph by Larry W. Richardson. B) Two orchid pollinia with 1 mm scale bar. Photograph by Lawrence E. Reeves.

Figure 3-2. Map of the four locations at the Florida Panther National Wildlife Refuge from which Encyclia tampensis flowers were sampled during the three-year study. Location 1 is the developed site, and locations 2-4 are the natural swamp habitat. Three replicates were utilized in each location.

Figure 3-3. A fine mesh exclusion bag placed around two Encyclia tampensis flowers that had opened soon after being covered. Exclusion bags were used to determine if insect pollinators were important to seed capsule production. These bags prevented any potential pollinators from visiting the flowers. Photograph by Haleigh A. Ray.

Figure 3-4. Flowers of Encyclia tampensis. A) Enlarged green receptacle of flowers indicate that the flowers have been pollinated. B) Another pollinated flower with enlarged receptacle. C) Slender, yellow receptacle of the lower flower shows no evidence that pollination has occurred. Photograph by Haleigh A. Ray.

Figure 3-5. Three types of insect traps were used to sample potential pollinators of Encyclia tampensis, the Florida butterfly orchid. A) Blue vane traps. B) WHY (wasp, hornet, yellow jacket) traps. C) Painted insect bowl traps. Photographs by Haleigh A. Ray.

Figure 3-6. Total number of insects actively sampled from each location at the Florida Panther National Wildlife Refuge where Encyclia tampensis was flowering over a three-year period (2015-2017). Location 1 had significantly more pollinators than did any of the other three locations (P = 0.0057), which were not significantly different from one another. Each of the points at the locations represent the insect orders Coleoptera, Hymenoptera, and Diptera, and the number of insects collected from each. Diamonds represent 95% confidence intervals for each mean.

Figure 3-7. Total number of insects in each order actively sampled from Encyclia tampensis flowers at the Florida Panther National Wildlife Refuge over a three-year period (2015-2017). There was no significant difference between the mean total number of insects in each order that were sampled (P = 0.4142).

Figure 3-8. Difference in average number of insects sampled around Encyclia tampensis flowers in the morning (0800-1200 h - AM) compared to the afternoon (1230-1630 h - PM) at four locations in the Florida Panther National Wildlife Refuge. A t-test was performed on the data and overall there was no significant difference in the number of insects collected in the morning compared to the afternoon at each site (t=0.877).

CHAPTER 4 EFFECT OF POLLINATION AND LOCATION ON SEED GERMINATION OF TWO NATIVE ORCHIDS

Introduction

In Florida, loss of hardwood hammock and swamp habitat has negatively impacted understory plant populations, such as native orchids (Sprott and Mazzotti 2001). These hardwood hammocks and swamps are prevalent at the Florida Panther National Wildlife Refuge (FPNWR), a 26,400 acre refuge located in Collier County, FL. This refuge is home to at least 27 species of native orchids found in these habitats (Stewart and Richardson 2008), and many of the orchids found there are listed as either threatened or endangered on Florida's Regulated Plant Index. The two orchids from this study are Prosthechea cochleata and Encyclia tampensis, which are listed as endangered and commercially exploited, respectively. Both of these orchids are epiphytic, and can be found in several swamp locations throughout FPNWR.

Orchid seeds are extremely small, with a length of 0.25-1.2 mm and width of 0.09-0.27 mm, and each capsule may have anywhere from 1,300 to 4,000,000 seeds (Knudson 1922,

Arditti 1967). Seed germination in orchids is unusual due to the fact that orchid seeds lack an endosperm. Without an endosperm, the seed must be infected with a mycorrhizal fungus to gain the nutrients necessary for development. There are two ways to grow orchid seeds in a lab.

Symbiotic germination, or giving the orchid seeds a fungus for germination, and asymbiotic germination, in which the seeds are grown in a Petri dish on a nutrient agar. For mass propagation of orchid seeds, asymbiotic germination has been successful (Kauth et al. 2008).

Several orchid species have been successfully propagated using these methods, including several species native to south Florida (Stenberg and Kane 1998, Johnson et al. 2007, Dutra et al. 2008,

Dutra et al. 2009a). When growing these seeds in the lab, they progress through several stages of

development, rated on a scale of 1-5 (Dutra et al. 2009a). The ability to grow large numbers of these orchids in a laboratory setting is an important aspect of orchid conservation.

Previous work on P. cochleata (known then as Encyclia cochleata) examined multiple populations of the species, some of which were self-pollinating and others which were not

(Ortiz-Barney and Ackerman 1999). While E. tampensis has been shown from Chapter 3 to be incapable of self-pollination, it is not known how readily the population of P. cochleata in south

Florida would self-pollinate, and if plants located in different types of habitats would self- pollinate at different rates or produce more viable seeds. In this study, P. cochleata and E. tampensis seed capsules (including any that are known to have self-pollinated) were collected from different areas in FPNWR and taken to Gainesville, FL for asymbiotic germination. The goal of this research was to determine any differences in seed capsule development or the percent of total seed germination from seed capsules collected from open, covered, or hand cross-pollinated flowers, or by location at FPNWR.

Materials and Methods

Prosthechea cochleata

The first step was to place fine mesh exclusion bags on some of the unopened flowers

(Figure 4-1). Over the course of two years at FPNWR, a total of 47 flowers were bagged out of

201 blooms, approximately 23%. Of these, 14 were from a developed location at FPNWR, and the remaining 33 from two separate locations that are natural habitat. In Gainesville, FL at the

University of Florida, potted P. cochleata flowers were kept in a greenhouse and had 29/56, bagged flowers, approximately 52%. In addition to these numbers, 12 hand cross-pollinations were performed in the greenhouse by moving the pollinia of one flower to the stigma of a flower on a different plant.

After counting and placing exclusion bags, the flowers could either senesce or produce a seed capsule. Any seed capsules formed were left to develop for 12-14 weeks before being collected and taken to a laboratory at the University of Florida in Gainesville. In the laboratory, they were processed as in Stewart and Kane (2006) and Dutra et al. (2008). The capsules were first placed into 50 mL plastic centrifuge tubes and kept over indicating silica gel desiccant until they began to dehisce (approximately two weeks). Once the capsules began to dehisce, they were labeled and stored in parafilm sealed centrifuge tubes in the freezer until being germinated.

For germinating the seeds, P723 orchid seed sowing medium from PhytoTechnology

Laboratories was used. The powder was dissolved into water at the suggested rate (32.74g/L) into a 500 mL Erlenmeyer flask, the lid covered with Reynolds Wrap® tin foil, and autoclaved for 40-55 minutes at approximately 121°C. In addition to the P723 media, two 100 mL empty beakers and another beaker with 90 mL of DI water was autoclaved at the same time. After being removed from the autoclave, everything was allowed to cool for 20 minutes. Once cooled, the medium was used to fill Petri dishes (8.5 mm diameter, 1.4 mm depth) approximately ½ full, and left to solidify. The 90 mL of DI water that was autoclaved was then mixed with 5 mL of 100% ethanol and 5 mL Clorox® concentrated bleach (8.25% sodium hypochlorite). This mixture was used to surface sterilize the seeds after removing them from the seed capsule. To surface sterilize the seeds, the mixture was added to a centrifuge tube with seeds and shaken for one minute. The mixture was removed, and sterile DI water added to rinse the seeds. The seeds were rinsed twice, then left in clean, sterile water. Approximately 0.1 mL of the seed solution was added to each of four sub-replicates on the Petri dish (Figure 4-2). For each seed capsule, at least eight plates with four sub-replicates were made, for a total of at least 32 replicates per capsule. After the seeds were dispersed, the Petri dishes were sealed with parafilm and stored at 24 ± 2ºC with a 14:10h

L:D cycle. These methods were based on previous work from Stewart and Kane (2006), Dutra et al. (2008), and Dutra et al. (2009b).

Encyclia tampensis

At FPNWR, there are several locations throughout the refuge where E. tampensis plants are present. While most of them are found in the undisturbed pond habitats, there are some plants growing at a developed, landscaped work center. Based on previous research, it has been determined that E. tampensis flowers are unable to self-pollinate, and require a pollen vector

(Chapter 3). Therefore, the only variable in seed capsules for this species was the location from which they were collected. After seed capsules were collected from each of the ponds and the work center, they were taken back to University of Florida to be processed, germinated, and stored in the same way as the P. cochleata capsules.

Statistical Methods

I analyzed the germination percentage of seeds from capsules of different treatments by using a one-way analysis of variance (ANOVA) with JMP 9.0.2 (SAS Institute 2010). The independent variables were the treatments (open, bagged, hand cross-pollinated), and the dependent variable was the germination percentage. A Tukey-Kramer test was used to separate means (α < 0.05) and determine any differences in percent germination between locations. To determine the percentage of germinated seeds, I divided the total number of germinated seeds by the total viable seeds in each of the sub-replicates and calculated the standard deviation.

Results

Prosthechea cochleata Seed Capsules

Over the course of the two year study, 43 of the 47 bagged flowers (91.5%) produced seed capsules. The four flowers that were bagged but did not form seed capsules were all at the developed location. A total of 10 of the 43 bagged capsules were collected, and an additional 10

capsules from flowers that were left open. Eight capsules were also collected from hand cross- pollinated flowers (pollinia from a flower, moved to a flower on a different plant), as that was all that developed. There were an average of 77.8 ± 37.7 seeds on each of the Petri dish sub- replicates from open capsules, and 76.5 ± 24.2 seeds per sub-replicates from bagged capsules.

However, there was an average of 173.7 ± 46.5 seeds on each sub-replicates from flowers that were hand cross-pollinated. Despite this difference, 37.8% of seeds from open flowers germinated, 41.6% from bagged, and 39.2% from hand cross-pollination (Figure 4-3). There was no significant difference between mean germination percentage from any of the capsule groups

(P=0.068)

When comparing germination data from each of the three locations at the FPNWR, there was no significant difference between the percent germination of seeds from capsules collected from the developed work center, the first natural area, or the second natural area (P=0.438)

(Figure 4-4).

In addition to the germination data presented above, it was observed and noted that the seeds from capsules that were hand cross-pollinated reached growth Stage 5 (Figure 4-5) in approximately four weeks, whereas the seeds from open and bagged FPNWR capsules took approximately five weeks to reach the same stage.

Encyclia tampensis Seed Capsules

Though no covered flowers produced seed capsules, 10 capsules from uncovered flowers were collected. There were four collected from the developed center, and three collected from each of the two natural habitats. Zero capsules developed from hand self-pollination, supporting the previous data that E. tampensis is not capable of self-pollination.

The average number of germinated seeds per sub-replicates were 10.3±4.3 at the work center, and 10.8±4.44 and 10.5±4.6 at the two natural habitat sites. This was from an average of

35.5±10.6, 35.4±11, and 35.8±13.2 total seeds per sub-replicates, respectively. For the percentage of germinating seeds, there was no significant difference between capsules collected from each of the locations (P=0.896) (Figure 4-6). There was 29.4% germination at Location 1,

29.9% at Location 2, and 28.9% at Location 3.

Discussion

The endangered and commercially-exploited status of P. cochleata and E. tampensis illustrate the need for more knowledge about their reproductive biology. In this study, I found that the P. cochleata seed capsules collected that were hand cross-pollinated and those that were either left open or bagged had about the same percentage of seeds germinate successfully.

However, the seed capsules from P. cochleata flowers that were hand cross-pollinated produced more seeds than those that were bagged or open at FPNWR. It's possible that even the open flowers at FPNWR are self-pollinating, causing there to be little difference in seed capsule formation and seed germination. From the results of the hand cross-pollination of greenhouse specimens, it suggests that these flowers may be producing seed capsules with more seeds if they are cross-pollinated, as opposed to self-pollinating. Hand self-pollination was not performed in the greenhouse as it has been performed in a previous study (Ortiz-Barney and Ackerman 1999), and the flowers at FPNWR were already known to likely be pollinating from the exclusion bag studies.

Ortiz-Barney and Ackerman (1999), examined inbreeding depression and the effect of hand self-pollinating and hand cross-pollinating P. cochleata (known then as Encyclia cochleata), and found that 34% of seeds from self-pollination were germinating, and only 24% from cross pollination. My percentages of seed germination were closely aligned, at 38.5% and

39.1% respectively.

Seed capsules from both P. cochleata and E. tampensis produced seeds with an average germination of 39.7% and 29.4%, respectively. With such a low percentage of seeds germinating on nutrient media, it is likely that an even smaller percentage will be germinating in nature.

However, being able to grow seedlings in the laboratory can be beneficial for conservation. The plants that develop from laboratory-grown seeds could be introduced into a natural area such as the FPNWR. With there being no significant difference between germination at the three locations from this study, it suggests that plants could be out planted at all areas of the FPNWR, and future seed capsules would have similar chances of producing viable seeds.

One goal of this research was to determine if there were any differences in seed capsule production in open flowers or those with exclusions bags. In addition, I wanted to see if there were any differences in the percent of seed germination from the three treatments (open, bagged, hand cross-pollinated), or across any of the locations at FPNWR from which the capsules were collected. I was able to determine that P. cochleata is capable of self-pollination, with seed capsules being produced in both open and bagged flowers. I found that while there was no significant difference between the percent of total seed germination between the capsules, the capsules that were hand cross-pollinated produced more seeds than those that were open or covered, both of which were likely self-pollinating.

Figure 4-1. Mesh exclusion bag placed on an unopened bloom of Prosthechea cochleata in the Florida Panther National Wildlife Refuge. Photograph by Haleigh A. Ray.

Figure 4-2. Example of four sub-replicates of Prosthechea cochleata seeds on P723 orchid seed sowing agar in a Petri dish. Photograph by Haleigh A. Ray.

Figure 4-3. Percent total germination of Prosthechea cochleata seed capsules that were covered with exclusion mesh (Bagged) at the Florida Panther National Wildlife Refuge (FPNWR), hand cross-pollinated (Hand Crossed), or left open at the FPNWR (Open). Percent germination was counted three to four weeks after seeds were plated onto the medium. Mean germination percentages with the same letter are not different by Tukey- Kramer test (α < 0.05).

Figure 4-4. Comparison of Prosthechea cochleata seed germination across three locations in the Florida Panther National Wildlife Refuge (FPNWR). There was no significant difference between the percentage of seed germination from capsules collected at each of these three locations (P=0.438).

Figure 4-5. Six Prosthechea cochleata seeds (Stage 5) among numerous other seeds that did not germinate. Photograph by Haleigh A. Ray.

Figure 4-6. The percentage of germinating Encyclia tampensis seeds from capsules in each of three locations at the Florida Panther National Wildlife Refuge. Location 1 is a developed work center, while Locations 2 and 3 are natural habitat. There is no significant difference between germination percent at these locations (P=0.896).

CHAPTER 5 DETERMINING POTENTIAL POLLINATORS OF Prosthechea cochleata (ORCHIDACEAE) AND ATTRACTION OF HONEY BEES TO A P. cochleata SYNTHETIC FLORAL FRAGRANCE BLEND

Introduction

Pollination is an essential aspect of flowering plant biology. In addition to flower colors, floral fragrance is another key component of pollinator attraction. Actively collecting flower- visiting insects from a flower can be a strong indicator of pollinator species, and many of these visitors are being attracted to the flower due to the fragrance. When collecting insects that are attracted to a flowers fragrance, there is another option for successful capture of pollinators besides active sampling.

Several types of traps have been used with synthetic fragrance blends to attract potential orchid pollinator species. McPhail traps have been commonly used when trapping orchid bees, or euglossine bees (Bennett 1972, Becker et al. 1991, Botsch et al. 2017). Blue vane traps are another type of trap that has been effective for catching native bees, including euglossine bees. In

Florida, they have been used to determine the range of Euglossa dilemma, an introduced euglossine bee (Pascarella 2017), and in Illinois they were used to assess bee diversity in prairie habitats (Geroff et al. 2014). Traps with fragrance blends that have been used for collection of other insects are moth buckets, Japanese beetle traps, and stink bug traps (Wellso and Fischer

1971, Mitchell et al. 1989, Chen et al. 2014, Guerrero et al. 2014, Sargent et al. 2014, Tillman et al. 2017).

Another type of experiment that utilizes fragrances and attraction by Hymenoptera is Y- tube olfactometer tests. A study by Brodmann et al. (2008) used Epipactis helleborine orchid fragrance to test wasp pollinators, placing them in Y-tubes with green leaf volatiles that were found to attract the wasps. A different orchid, Dendrobium sinense, was found to produce

volatiles that attract prey-hunting hornets, as it mimics bee alarm pheromones (Brodmann et al.

2009). Y-tubes were also used to show that the hornets were attracted to the flower volatiles.

Prosthechea cochleata is a state endangered, epiphytic orchid that can be found in south

Florida. One study in Mexico determined that the flowers did not produce any floral volatiles

(Cancino and Damon 2006), but an experiment on these flowers in south Florida found that they were producing a fragrance (Ray et al. 2018). Synthetic floral blends can be created based on compounds identified from GC/MS floral fragrance analysis. In this study, I first identify potential pollinators of P. cochleata by actively sampling insects visiting the flowers as well as using traps. Then, I use a fragrance blend created from volatile compounds identified in P. cochleata flowers. This floral blend was used to gather preliminary data on honey bee (Apis mellifera) attraction to the synthetic odor. The fragrance was used in baited traps in four different locations as well as in Y-tube experiments. The goal of this research was to gather preliminary data if bees, specifically honey bees, were attracted to the floral odor I produced to mimic the odor emitted by P. cochleata.

Methods

Pollinator Sampling

Three locations at FPNWR where P. cochleata flowers can be found were used for the pollinator sampling. Potential pollinators were collected using both active sampling and traps.

When active sampling, blooming flowers at each location were monitored for 45 minutes in the morning, and 45 minutes in the afternoon for 90 total minutes of observation per location each day, for five days each year (2015 and 2016). Any collected specimens were preserved and taken to the University of Florida Entomology and Nematology Department for identification.

The traps that were used in combination with active sampling were Blue vane traps

(SpringStar ®, Seattle, WA), W.H.Y. (wasp, hornet, yellow jacket) traps (Rescue ®, Spokane,

WA), and insect bowl traps comprised of three differently colored bowls: yellow, white, and blue

(Blue Sky®, Brooklyn, NY). Three of each trap were set up at each of the three locations at

FPNWR, all within 4-5 meters from flowering orchids. Each trap contained water with approximately 0.01% Silwet L-77 surfactant (Helena Chemicals ®, Collierville, TN) to prevent insect escape. While the bowl traps were placed at ground level, the other two traps were suspended about 1.5 m high. All traps were checked daily after the afternoon active sampling, and trapped insects removed and preserved for identification.

Floral Odor

Based on previous experiments (Chapter 2), it was determined that Benzaldehyde,

Mesitylene, Nonanal, Decanal, Ocimene, Pseudocumene, Pinene, and Octanal were the most abundant floral fragrance compounds in P. cochleata. Chemical compounds of these (Sigma

Aldrich ®) used to create the floral blend were stored at the USDA in Gainesville, Florida. The individual floral compounds were added to pentane. The pentane evaporated, leaving only the floral blend that I then used to determine bee attraction. The compounds were diluted to 1 μl /1 ml of pentane before being added at a ratio similar to the amounts found in the P. cochleata fragrance (Table 5-1). Once the blend was created, 1 ml was added to microcentrifuge tubes containing a 3-4 cm piece of cotton wick. The wick absorbed most of the blend, leaving a small amount pooled in the bottom of the tube. A 2 mm hole was placed in the microcentrifuge tubes lid to allow the odor to disperse. A single tube containing the odor was placed into each trap, additional traps contained a blank control. The same was done for the Y-tube experiments.

Insect Trapping

Six traps were used and set up at the USDA honey bee apiary to screen trap types for effectiveness. The traps used were: blue vane traps (SpringStar ®, Seattle, WA), W.H.Y. (wasp, hornet, yellow jacket) traps (Rescue ®, Spokane, WA), McPhail traps (Great Lakes IPM ®,

Vestaburg, Michigan), stink bug traps (Rescue ®, Spokane, WA), Japanese beetle traps

(Tanglefoot ®, Marysville OH), and moth bucket traps (Uni-Trap, distributed by Great Lakes

IPM ®, Vestaburg, Michigan). Traps were suspended approximately 1.75 m from the ground on a metal plant shepherd’s hook (Garden Treasures, North Wilkesboro, NC). Each trap contained one microcentrifuge tube with 1 ml of the floral blend, and was filled approximately 1/3 of the way full with water and the surfactant Silwet L-77 (Helena Chemical Company, Collierville,

Tennessee). The traps were checked twice daily for four days, after which the floral blend was likely to no longer be as attractive. Therefore, I was able to determine trap effectiveness for collecting Hymenoptera, and have a better understanding of when the bees were most active.

The two most effective traps, Trap A and Trap C (blue vane and moth bucket traps), were then set up as previously stated at four study sites: USDA-ARS-CMAVE, Gainesville, FL

(USDA); Natural Area Teaching Lab (NATL) on the University of Florida (UF) campus; UF

Honey Bee Research and Extension laboratory (HBREC); and a private, rural, wooded residence located in Alachua, Florida. In addition to the two traps, a control trap for each trap type was also used at each location. Traps were checked twice daily, between 0600 and 0800 h, and between

1600 and 1800 h, for four consecutive days in November 2017. The experiment was repeated every week for four weeks. Specimens collected were taken to the UF Entomology and

Nematology Department for identification.

Y-tube Behavior

The floral blend created for the insect trapping was used in Y-tube olfactometer tests. In this experiment, I tested both the lab created floral fragrance, as well as live P. cochleata flowers. A single flower (attached to the plant) or the floral blend was placed in a Reynolds ® oven bag at one arm of the tube, and the other arm was left as a blank control. Filtered air was pumped through the bags and into the arms of the Y-tube. An additional pump was set up to pull

the air from the Y-tube, providing airflow through the tube (Figure 5-1). The Y-tube was set in a fume hood, and covered with a light-proof cloth to limit visual cues and light attraction that might affect A. mellifera decisions. Two separate Y-tubes were used, one for the flower and one for the volatile blend, thus preventing cross contamination between fragrances.

Apis mellifera collected from hives at the USDA in Gainesville, Florida were used in the

Y-tubes to determine if they displayed attraction to the orchid flower fragrance. Bees were collected from the hives approximately 30 minutes prior to starting the experiment. They were kept in insect cages with access to water and honey. A single honey bee, collected into a small plastic vial was released into the end of the Y-tube and given five minutes to make a choice. If the bee moved down the tube, and at least 6 cm into an arm (approximately 1/3 of the arm length), the time it took to make the choice was recorded along with whether they chose the fragrance or the control arm. This was repeated 20 times for the flower and 20 times for the fragrance blend, using a new bee for each time, for a total of 40 replicates per day. After ten replicates, the Y-tube was flipped, while continuing to pull the fragrance through the same arm, to rule out any contamination issues or potential positional bias, such as light cues that the bees were attracted to or possible contamination by lingering volatiles. This was repeated over the course of three days, for a total of 120 replicates.

Statistics

For statistical analysis of this preliminary Y-tube data, I used the Wilcoxon Signed Rank test to compare the selections made by the bees, either the fragrance arm of the tube or the control arm of the tube. A t-test was performed to test for any difference in pollinator abundance in the morning collections compared to the afternoon collections.

Results

Pollinator Sampling

Over the course of the study, only 11 insects were actively sampled from P. cochleata flowers. All were in the Hymenopteran family Apidae. Six were Euglossa dilemma, the non- native orchid bee found in south Florida, three were from the genus Melissodes, and two from the genus Bombus. Of these 11, zero were collected from the developed location, all were from the natural habitats. In addition to active sampling, there were 37 insects collected in the traps.

Only four of these were in the family Apidae, with the others typically being small Diptera.

Insect Trapping

Overall, few insects were collected in the blue vane and moth bucket traps over the sampling period. A total of 14 Hymenoptera (not including Formicidae) were collected from fragrance traps, 11 in the vane traps and three in the bucket traps. There were eight Hymenoptera collected from control traps, six of which were A. mellifera. Of the total number of Hymenoptera captured (Appendix B) (fragrance and control), 16 were A. mellifera, which were only collected from USDA and HBREC. There were more Hymenoptera collected when checked in the afternoon (16) compared to when checked in the morning (6), but there was not a significant difference between the two (P = 0.246). The only other insect orders that were collected were

Diptera (18) and Coleoptera (2).

Y-tube Behavior

When placed in the Y-tube, the bees chose the fragrance side, either the orchid flower or the volatile blend, 50 times (41.67%), compared to choosing the control side 44 times (36.67%).

During 26 of trials (21.67%), there was no choice made by the bees. There was no significant difference between the number of times that a bee chose the fragrance or control, when using either the volatile or the flower (P = 0.250). Additionally, there was no difference between the

number of times the bees chose the fragrance or the control when using the flower compared to the volatile. During the trials using the flower, bees chose the fragrant side 25 times and the control 22 times (13 no choice), and when using the volatile, bees again chose the fragrant side

25 times and the control 22 times (13 no choice). When exposed to either volatile blend or control, the bees that chose the blend did so in approximately 143.6 ± 58.4 seconds, compared to those choosing the blank in 139.9 ± 70.1 s. When given the option of the actual flower, those that chose the flower did so in 133.5 ± 56.2 s, or chose the blank in 157.4 ± 77.5 s.

Discussion

Overall, there was very low capture of trapped insects, including A. mellifera. Of all four locations, the USDA site had the highest capture, with the Alachua site having the lowest. One possible explanation for USDA having the highest capture was the proximity of the traps to honey bee hives. Though another site was the UF Honey Bee Research and Education Center, the traps were placed further from hives there (approximately 20-25 meters) than at the USDA

(approximately 4-5 meters). Also, not all Hymenoptera from the USDA site were A. mellifera.

There were five additional Hymenoptera captured as well. However, based on the insect sampling at FPNWR, A. mellifera is unlikely to be pollinating P. cochleata. This could be because the flower does not provide an attractive enough fragrance or food source for the honey bees, or because the bees are not as frequently seen at FPNWR as they are at the University of

Florida. Previous studies have suggested that honey bees could present enough competition to native bees to negatively affect native bee populations (Dupont et al. 2004, Thomson 2004).

Based on this, it would have been expected that the native bees would be more abundant in the

Alachua and NATL locations, and not frequently collected at the USDA or Honey Bee Research and Education Center locations.

Another difference noted was that the blue vane traps collected more Hymenoptera than did the moth bucket traps. It has been shown through behavior tests that honey bees prefer blue

(Giurfa et al. 1995, Morawetz et al. 2013, Dyer et al. 2007). This makes it likely that the difference in trap capture could be attributed to trap color. When comparing the traps with the floral blend to control traps, there was not a significant difference in the number of

Hymenoptera captured. When replicated in the future, it may be equally as effective to use only the blue vane traps, or to set up both traps at different times so that color is not a factor.

For the Y-tube experiment, bees chose the fragrant side over the control the exact same number of times when using the flower or the volatile blend, suggesting there was no different between attraction to the flower compared to attraction to the blend. Even though the blend seemed to be equally as attractive as the flower, there was still not a significant difference in choice between the fragrant side or the control side. In previous pollinator work on P. cochleata, only bees of the family Apidae were collected (Chapter 4). However, none of the bees collected or seen visiting the flowers of P. cochleata were A. mellifera. This could mean that because the bees did not have a preference for the flower, that they would not be pollinators of the flower in the wild, or that any pollinating bees are not in the area.

Future research may be directed at field studies investigating attraction and pollination to the orchid flowers by A. mellifera. Conducting the trapping experiments during P. cochleata blooming season in the orchids native habitat would need to be performed. It is possible that A. mellifera are not potential pollinators of P. cochleata, and that another native bee with a more restricted range is an effective pollinator of P. cochleata. Trapping with the floral fragrance blend near blooming flowers would increase the chances of collecting insects with orchid pollinia attached, thus verifying it as a pollinator. If another species was abundantly collected,

the Y-tube tests could potentially be repeated using that species. Apis mellifera is a widespread pollinator and may have the potential to pollinate this state endangered orchid along with other native pollinators.

Table 5-1. Eight compounds created the floral blend that was in each of the six traps. The amount listed in the right column is the amount of microliters per milliliter of each compound dilution in pentane that was used. Compound Concentration Octanal 150 μl Mesitylene 100 μl Benzaldehyde 100 μl Ocimene 50 μl Nonanal 20 μl Decanal 20 μl Pseudocumene 20 μl Pinene 20 μl

Figure 5-1. Y-tube olfactometer set up in the laboratory. A screen was kept over the main part of the tube to reduce visual cues for Apis mellifera. A) Side of tube with an empty attached bag to serve as a control. B) Side with a microcentrifuge tube containing orchid fragrance volatiles. Photograph by Haleigh A. Ray.

CHAPTER 6 CONCLUSIONS AND FUTURE WORK

Pollination of orchids is highly variable across species, leaving much to study regarding reproduction in this large plant family. Some orchids may use deception (chemical or physical, typically sexual or pheromone based), and they can have a very specific relationship with a single pollinator, or be generalist pollinated by mammals, birds, or invertebrates. The pollination biology of the orchid species I focused on in these studies, Encyclia tampensis and Prosthechea cochleata, had not been previously studied in depth. The research from this dissertation was conducted to give a more comprehensive understanding of their reproductive biology. The goals of this research were to determine the floral fragrance of P. cochleata, identify potential pollinators of both P. cochleata and E. tampensis, find out whether or not these species are capable of self-pollination, and to compare the resulting seed capsules and seed germination of both P. cochleata and E. tampensis. I have also begun collecting preliminary data on the attraction of pollinators using synthetic floral fragrance blends.

In summary, in the first part of my research the floral fragrance of P. cochleata yielded results which conflict with previous results to what I was seeing (or rather smelling) in the

University of Florida greenhouses. Though it was previously reported that P. cochleata did not produce a fragrance, volatiles were collected from the flowers growing in the greenhouse and in the wild in south Florida at the Florida Panther National Wildlife Refuge (FPNWR). All of these volatiles had been previously recorded in other flowering plant species, many from the family

Orchidaceae. These volatiles were then used to create a fragrance blend for preliminary testing with baited trapping and Y-tube experiments.

The second part of this research involved trapping and actively collecting potential pollinators from FPNWR when E. tampensis was in bloom. This also included placing exclusion

bags over flowers to determine if they were capable of autogamy (self-pollination). Six families of insects within three orders (Coleoptera, Diptera, Hymenoptera) were actively collected from flowers during E. tampensis blooming, suggesting that this species is generalist pollinated. To further the idea that E. tampensis is using pollen vectors, none of the 231 flowers covered with exclusion bags developed seed capsules. Based on this data, it is highly probable that E. tampensis is not capable of self-pollination.

The third part of this dissertation was to germinate orchid seeds from P. cochleata and E. tampensis seed capsules that developed from both cross- and self-pollination. This allowed me to determine if capsules resulting from cross-pollination produced more seeds or seeds with a higher percentage of germination than capsules produced from self-pollinated plants, or if there was no difference in seed germination. While E. tampensis is not able to self-pollinate, over 90% of P. cochleata flowers produced seed capsules inside of the exclusion bags.

After completing the first three parts of this dissertation research, the pollination biology of P. cochleata still was not clear. Even though they were producing a fragrance, the flowers seem to be readily self-pollinating in nature. However, when the resulting seeds were compared, cross-pollinated flowers were producing capsules with more seeds. This led to another set of experiments involving a laboratory created fragrance blend that mimics the fragrance of P. cochleata. During P. cochleata blooming, the same trapping methods from the previous study were done in an attempt to identify potential pollinators of P. cochleata. Only one family of insects was actively collected from the flowers (Hymenoptera: Apidae).

The first part of this was to put the blend into two traps, and place them in areas that bees would be likely to visit. Traps were chosen based on preliminary data using six traps, then the two most effective traps were used for further experiments. Few Hymenoptera were collected

over the course of the study, but were most abundant in the blue vane traps baited with my volatile blend.

Honey bees were used in Y-tube experiments in the laboratory. One side of the tube was a control, and the other used either the laboratory created blend, or a blooming P. cochleata flower. There was no significant difference in which side of the Y-tube the honey bees selected.

The results from these two projects indicate that honey bees may not be attracted to the fragrance of P. cochleata. Further tests are planned to continue this research in the future.

In addition to this work, another project was started, but I was unable to complete the project due to technical difficulties. I attempted to use video cameras to record pollinator activity on the orchids, similar to the projects in Steen (2012), Steen and Aase (2011), and Suetsugu and

Hayamizu (2014). I used a waterproof, IR, CCD camera usable for day or night recording, a mini digital video recorder (DVR) system, and a 12-volt sealed lead battery. The battery and the DVR system were kept in a sealed plastic tote to prevent water damage to the equipment. All video was recorded on a 4.0 GB SanDisk memory card. The products were chosen based on the equipment used in Steen and Aase 2011.

However, when attempting to set up the motion detection component, I found that the equipment I was using was capturing movement from wind moving the plants, causing it to continuously record. Additionally, since both E. tampensis and P. cochleata are both epiphytic, the cameras would have had to be suspended somehow. Because I was using a large battery to power the camera traps, this didn't allow me to be able to hang the traps in any safe way.

One other obstacle for camera use on P. cochleata was that this species blooms during the rainy season at my field site. Typically, I have recorded 1-1.5 m of standing water around all flowering plants. This creates an unstable floor to elevate camera traps from the ground.

Overall, I believe that camera traps can be extremely useful in identifying pollinating species of flower-visiting insects, including orchid species. The orchids used in the camera trapping studies above (Steen 2012, Suetsugu and Hayamizu 2014) were from the genus

Platanthera, which are terrestrial species, not epiphytes such as in my study. One way to set up this project differently would be to use a smaller battery. This would allow the traps to be mounted to a tree branch more easily, getting the cameras much closer to the orchid flowers. The downside of this would be that the batteries would need to be charged more frequently, and couldn't be left running for multiple consecutive days. Additionally, a motion detection software such as Zone Trigger could be used. With this, the video could continuously record and the software detects motion inside of a "hot spot," saving the video during the times there is movement in that spot.

In addition to the projects detailed in this dissertation, I'm beginning a comprehensive review of orchid pollination. This has begun with a literature review to determine all orchid species with known pollinators. I will collect the identification of the pollinators, and the location of the orchid. Once complete, the R package 'bipartite' will be used for creating a pollinator web and for any statistical analysis. This package is used to visualize and describe patterns in ecological webs consisting of two levels, such as orchid species and their pollinators. Currently, I have data from 47 orchid species, with pollinators from 19 insect families. After completing data collection about insect pollinators, I plan to also gather data from non-insect pollinators, such as birds and bats. The goal is for this information to be useful for all future work in orchid pollination, as well as potentially providing insight into pollinator patterns and likeliness of a species to be a pollinator of orchids.

As a small example for this pollinator web project, I have created a web using at least six species from three terrestrial orchid genera (Caladenia, Epipactis, and Spiranthes) (Figure 6-1).

From the 20 species used, there are pollinators from 12 insect families, eight Hymenopteran and four Dipteran.

Aside from this project, there are additional directions that future research on E. tampensis and P. cochleata could be conducted. From the floral fragrance work in Chapter 2, I determined that P. cochleata produced a fragrance at the same location, during blooming. It would be advantageous to collect volatiles from this species at additional locations, to see if/where there is a gradient that P. cochleata becomes less or non-fragrant. Because some flower species have variations in fragrance based on geographic location, this could be a factor affecting

P. cochleata, and the differences seen between my research and the study by Cancino and

Damon (2006). Another fragrance component that I would be interested in would be determining if the orchid produces less or no fragrance after being cross-pollinated, and also continuing fragrance collection until seed capsule formation in the case of self-pollinating flowers. Once a flower has been pollinated, it would seem to no longer have a known biological reason to continue producing fragrance. However, if P. cochleata is truly self-pollinating, it would be interesting to see when the fragrance production ceases.

Another way to further some of the research from this dissertation would be to verify the pollinators of E. tampensis. Though several of the species were actively collected from the flowers multiple times, collecting insects with pollinia attached would be the ideal way to confirm pollinator species. Many more hours of observation and trapping would likely be required, but if insects could be collected with pollinia, the pollinia on the insect could be compared to a known pollinia sample from the blooming orchids using visual techniques or

molecular techniques. It should be possible to determine the fragrance volatiles from E. tampensis like was done in Chapter 2 for P. cochleata, and potentially use a synthetic blend in traps, so that a more accurate representation of potential pollinators could possibly be collected.

One other option to more thoroughly sample for pollinators would be to include night sampling, to rule out the possibility of nocturnal pollinators visiting the flowers.

This type of research could be applied to other orchid species whose pollination biology is unknown or not fully understood. For example, there are several species at the Florida Panther

National Wildlife Refuge of which the pollination biology is unexplained. The ghost orchid,

Dendrophylax lindenii, is assumed to be pollinated by a hawk moth (Lepidoptera: Sphingidae), but this has not been confirmed by any research. The rigid epidendrum, Epidendrum rigidum, is assumed to be autogamous, but this is also unconfirmed. Johnson et al. (2009) studied the reproductive biology of Eulophia alta, a native Florida orchid thought to be autogamous, and found that autogamy occurred in less than 8% of flowers. It is possible that other species thought to be autogamous may be capable of it, but autogamy may not be the primary method of pollination.

The future of research on the pollination of E. tampensis and P. cochleata, as well as other orchid species, has numerous ways to be further explored. The research from this dissertation provides some of the building blocks to expand our knowledge of orchid pollination, including floral fragrance of the flower, the pollinators that it may attract, whether the orchids need those pollinators or if they are capable of self-pollination, and the viability of the resulting seed capsules. Combining this information can promote future conservation efforts for these orchids in an endeavor to rebuild or reestablish their populations.

Figure 6-1. Preliminary results of an ecological web created with the R package 'bipartite' comparing orchid species to their identified pollinator. This data compares species from three terrestrial orchid genera (Caladenia, Epipactis, and Spiranthes) to the insect family that pollinates them (Hymenoptera in red, Diptera in yellow).

APPENDIX A INSECTS COLLECTED DURING Encyclia tampensis BLOOMING

Table A-1. Insects collected from the Florida Panther National Wildlife Refuge during Encyclia tampensis blooming from 2015-2017. Order Family Genus Species Collection AM/PM Loc 1 Loc 2 Loc 3 Loc 4 Year Coleoptera Cantharidae Blue Vane n/a X 2015 Coleoptera Chrysomelidae Chrysomela scripta White Pan n/a X 2016 Coleoptera Elateridae Blue Vane n/a X 2015 Coleoptera Scarabaeidae Callistethus marginatus Blue Vane n/a X 2016 Coleoptera Scarabaeidae Callistethus marginatus Blue Vane n/a X 2017 Coleoptera Scarabaeidae Trigonopeltastes delta Active AM X 2015 Coleoptera Scarabaeidae Trigonopeltastes delta Active AM X 2015 Coleoptera Scarabaeidae Trigonopeltastes delta Active AM X 2016 Coleoptera Scarabaeidae Trigonopeltastes delta Active AM X 2016 Coleoptera Scarabaeidae Trigonopeltastes delta Active AM X 2017 Coleoptera Scarabaeidae Trigonopeltastes delta Active PM X 2015 Coleoptera Scarabaeidae Trigonopeltastes delta Active PM X 2015 Coleoptera Scarabaeidae Trigonopeltastes delta Active PM X 2017 Coleoptera Scarabaeidae Trigonopeltastes delta Blue Vane n/a X 2016 Diptera Dolichopodidae Blue Pan n/a X 2015 Diptera Dolichopodidae Blue Pan n/a X 2015 Diptera Dolichopodidae Blue Pan n/a X 2015 Diptera Dolichopodidae Blue Pan n/a X 2016 Diptera Dolichopodidae Blue Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2015 Diptera Dolichopodidae White Pan n/a X 2015 Diptera Dolichopodidae White Pan n/a X 2015 Diptera Dolichopodidae White Pan n/a X 2016 Diptera Dolichopodidae White Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2017

Table A-1. Continued Order Family Genus Species Collection AM/PM Loc 1 Loc 2 Loc 3 Loc 4 Year Diptera Dolichopodidae White Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2016 Diptera Dolichopodidae White Pan n/a X 2016 Diptera Dolichopodidae White Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2015 Diptera Dolichopodidae White Pan n/a X 2016 Diptera Dolichopodidae White Pan n/a X 2016 Diptera Dolichopodidae White Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2015 Diptera Dolichopodidae White Pan n/a X 2017 Diptera Dolichopodidae White Pan n/a X 2017 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2016 Yellow Diptera Dolichopodidae Pan n/a X 2017 Yellow Diptera Dolichopodidae Pan n/a X 2017 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2016 Yellow Diptera Dolichopodidae Pan n/a X 2016 Yellow Diptera Dolichopodidae Pan n/a X 2016 Yellow Diptera Dolichopodidae Pan n/a X 2016

Table A-1. Continued Order Family Genus Species Collection AM/PM Loc 1 Loc 2 Loc 3 Loc 4 Year Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2017 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2015 Yellow Diptera Dolichopodidae Pan n/a X 2016 Yellow Diptera Dolichopodidae Pan n/a X 2016 Yellow Diptera Dolichopodidae Pan n/a X 2017 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2015 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2015 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2016 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2016 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2016 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2017 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2016 Diptera Syrphidae Copestylum sexmaculatum Active AM X 2015 Diptera Syrphidae Copestylum sexmaculatum Active PM X 2017 Diptera Syrphidae Copestylum sexmaculatum Active PM X 2016 Diptera Syrphidae Copestylum sexmaculatum Active PM X 2016 Diptera Syrphidae Copestylum sexmaculatum Active PM X 2017 Diptera Syrphidae Copestylum sexmaculatum Active PM X 2016

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Table A-1. Continued Order Family Genus Species Collection AM/PM Loc 1 Loc 2 Loc 3 Loc 4 Year Diptera Syrphidae Copestylum sexmaculatum Active PM X 2017 Hemiptera Cicadidae Blue Pan n/a X 2015 Hymenoptera Apidae Bombus griseocollis Active AM X 2015 Hymenoptera Apidae Bombus griseocollis Active PM X 2016 Hymenoptera Apidae Bombus griseocollis Active PM X 2016 Hymenoptera Apidae Bombus pensylvanicus Active AM X 2015 Hymenoptera Apidae Bombus pensylvanicus Active AM X 2017 Hymenoptera Apidae Bombus pensylvanicus Active PM X 2015 Hymenoptera Apidae Bombus pensylvanicus Active PM X 2017 Hymenoptera Apidae Bombus pensylvanicus Blue Vane n/a X 2015 Hymenoptera Apidae Bombus pensylvanicus Blue Vane n/a X 2016 Hymenoptera Apidae Bombus pensylvanicus Blue Vane n/a X 2016 Hymenoptera Apidae Bombus pensylvanicus Blue Vane n/a X 2016 Hymenoptera Apidae Bombus pensylvanicus Blue Vane n/a X 2017 Yellow Hymenoptera Apidae Bombus pensylvanicus Pan n/a X 2017 Yellow Hymenoptera Apidae Bombus pensylvanicus Pan n/a X 2015 Hymenoptera Apidae Euglossa dilemma Blue Vane n/a X 2016 Hymenoptera Apidae Euglossa dilemma Blue Vane n/a X 2016 Hymenoptera Apidae Euglossa dilemma Blue Vane n/a X 2017 Hymenoptera Apidae Euglossa dilemma Blue Vane n/a X 2016 Hymenoptera Apidae Xylocopa micans Active PM X 2016 Hymenoptera Apidae Xylocopa micans Active PM X 2017 Hymenoptera Apidae Mellisodes bimaculata Active AM X 2015 Hymenoptera Apidae Mellisodes bimaculata Active AM X 2015 Hymenoptera Apidae Mellisodes bimaculata Active AM X 2016 Hymenoptera Apidae Mellisodes bimaculata Active PM X 2017 Hymenoptera Apidae Mellisodes bimaculata Active PM X 2017

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Table A-1. Continued Order Family Genus Species Collection AM/PM Loc 1 Loc 2 Loc 3 Loc 4 Year Hymenoptera Apidae Mellisodes bimaculata Blue Vane n/a X 2015 Hymenoptera Apidae Mellisodes bimaculata Blue Vane n/a X 2015 Hymenoptera Apidae Mellisodes bimaculata Blue Vane n/a X 2016 Hymenoptera Apidae Mellisodes bimaculata Blue Vane n/a X 2017 Hymenoptera Apidae Mellisodes bimaculata Blue Vane n/a X 2016 Hymenoptera Apidae Mellisodes bimaculata Blue Vane n/a X 2017 Hymenoptera Formicidae Pseudomyrmex gracilis White Pan n/a X 2015 Hymenoptera Formicidae Pseudomyrmex gracilis White Pan n/a X 2015 Hymenoptera Formicidae Pseudomyrmex gracilis White Pan n/a X 2015 Hymenoptera Formicidae Pseudomyrmex gracilis White Pan n/a X 2016 Yellow Hymenoptera Formicidae Pseudomyrmex gracilis Pan n/a X 2015 Yellow Hymenoptera Formicidae Pseudomyrmex gracilis Pan n/a X 2016 Yellow Hymenoptera Formicidae Pseudomyrmex gracilis Pan n/a X 2016 Yellow Hymenoptera Formicidae Pseudomyrmex gracilis Pan n/a X 2017

Hymenoptera Formicidae Pseudomyrmex gracilis Yellow Pan n/a 2017 Yellow Hymenoptera Formicidae Pseudomyrmex gracilis Pan n/a X 2017 Hymenoptera Halictidae Agapostemon Active AM X 2015 Hymenoptera Halictidae Agapostemon Active PM X 2016 Hymenoptera Halictidae Agapostemon Active PM X 2017 Hymenoptera Halictidae Agapostemon Active PM X 2017 Hymenoptera Megachilidae Osmia Active AM X 2015 Hymenoptera Megachilidae Osmia Blue Vane n/a X 2015 Hymenoptera Megachilidae Osmia Blue Vane n/a X 2016 Hymenoptera Megachilidae Osmia Blue Vane n/a X 2016

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Table A-1. Continued Order Family Genus Species Collection AM/PM Loc 1 Loc 2 Loc 3 Loc 4 Year Hymenoptera Vespidae Mischocyttarus Active AM X 2015 Hymenoptera Vespidae Mischocyttarus Active AM X 2016 Hymenoptera Vespidae Mischocyttarus Active AM X 2016 Hymenoptera Vespidae Mischocyttarus Active PM X 2017 Hymenoptera Vespidae Mischocyttarus Active PM X 2017 Hymenoptera Vespidae Vespula W.H.Y. n/a X 2015 Hymenoptera Vespidae Vespula W.H.Y. n/a X 2015 Hymenoptera Vespidae Vespula W.H.Y. n/a X 2016 Lepidoptera Hesperiidae White Pan n/a X 2016 Lepidoptera Hesperiidae White Pan n/a X 2016

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APPENDIX B INSECTS COLLECTED FROM TRAPS CONTAINING SYNTHETIC ORCHID FRAGRANCE

Table B-1. Insects collected from traps containing synthetic orchid fragrance. Preliminary data collected in 2017. Order Family Genus Species Collection AM/PM USDA NATL HBREC H.S. Hymenoptera Apidae Apis mellifera Blue Vane-F AM X Hymenoptera Apidae Apis mellifera Blue Vane-F AM X Hymenoptera Apidae Apis mellifera Blue Vane-F PM X Hymenoptera Apidae Apis mellifera Blue Vane-F PM X Hymenoptera Apidae Apis mellifera Blue Vane-F PM X Hymenoptera Apidae Apis mellifera Blue Vane-F PM X Hymenoptera Apidae Apis mellifera Blue Vane-F PM X Hymenoptera Apidae Apis mellifera Moth Bucket- F PM X Hymenoptera Apidae Apis mellifera Blue Vane- F AM X Hymenoptera Apidae Apis mellifera Blue Vane- F PM X Hymenoptera Halictidae Moth Bucket- F PM Hymenoptera Halictidae Blue Vane- F AM X Hymenoptera Thynnidae Blue Vane- F PM X Hymenoptera Scoliidae Moth Bucket- F PM Hymenoptera Apidae Apis mellifera Blue Vane-C AM X Hymenoptera Apidae Apis mellifera Blue Vane-C PM X Hymenoptera Apidae Apis mellifera Blue Vane-C PM X Hymenoptera Apidae Apis mellifera Moth Bucket- C PM X Hymenoptera Apidae Apis mellifera Blue Vane-C PM X Hymenoptera Apidae Apis mellifera Moth Bucket- C PM X Hymenoptera Halictidae Blue Vane- C PM X Hymenoptera Thynnidae Moth Bucket- C AM X

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BIOGRAPHICAL SKETCH

Haleigh Amanda Ray was born in 1990 in Springfield, Illinois, and grew up in Havana,

Illinois. She graduated from Havana High School in 2008 and went on to attend Illinois College, where she obtained her undergraduate degree in biology. While studying for her degree, she enrolled in an entomology course, and began working in the Orchid Recovery Program under the guidance of Dr. Lawrence W. Zettler. During this time she began work as an intern at the Florida

Panther National Wildlife Refuge, in Collier County, Florida, surveying native orchids for scale insect infestation. She became even more interested in orchids, leading her to pursue her master's degree in the Entomology and Nematology Department at the University of Florida in August

2012, working with Dr. Marjorie A. Hoy, which she earned in May 2014. At this time she studied a predatory mite discovered in association with arthropod pests of orchids, finding that it could feed on populations of three greenhouse pests. Wanting to continue her studies on orchids, she began a Ph.D. at the University of Florida in August 2014 with Dr. Jennifer L. Gillett-

Kaufman. Her research included in this dissertation focused on the pollination biology of two orchid species native to south Florida.

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